- Open Access
The population genetics of parasitic nematodes of wild animals
Parasites & Vectors volume 11, Article number: 590 (2018)
- The Correction to this article has been published in Parasites & Vectors 2019 12:468
Parasitic nematodes are highly diverse and common, infecting virtually all animal species, and the importance of their roles in natural ecosystems is increasingly becoming apparent. How genes flow within and among populations of these parasites - their population genetics - has profound implications for the epidemiology of host infection and disease, and for the response of parasite populations to selection pressures. The population genetics of nematode parasites of wild animals may have consequences for host conservation, or influence the risk of zoonotic disease. Host movement has long been recognised as an important determinant of parasitic nematode population genetic structure, and recent research has also highlighted the importance of nematode life histories, environmental conditions, and other aspects of host ecology. Commonly, factors influencing parasitic nematode population genetics have been studied in isolation, such that an integrated view of the drivers of population genetic structure of parasitic nematodes is still lacking. Here, we seek to provide a comprehensive, broad, and integrative picture of these factors in parasitic nematodes of wild animals that will be a useful resource for investigators studying non-model parasitic nematodes in natural ecosystems. Increasingly, new methods of analysing the population genetics of nematodes are becoming available, and we consider the opportunities that these afford in resolving hitherto inaccessible questions of the population genetics of these important animals.
Parasitic nematode infection is ubiquitous in wild animals and can profoundly alter the physiology, behaviour and reproductive success of hosts [1, 2], and as such parasitic nematodes play key roles in ecosystem functioning [3, 4]. However, parasitic nematodes can raise conservation concerns - invasive parasitic nematodes may threaten naïve, native hosts , while environmental changes may render hosts more susceptible to pre-existing parasitic nematode species, resulting in more severe disease . Furthermore, parasitic nematodes that normally infect wild animals can spill-over into human and domestic animal populations, acting as new sources of disease [7, 8].
Population genetic structure - the distribution of genetic variation in time and space - affects how a species responds to selection pressures, and so shapes its evolution . Studying the population genetics of parasites in wild animals has several benefits. First, it provides an insight into the parasite’s infection dynamics [9,10,11], with consequences for ecosystem functioning [3, 4]. Secondly, population genetic studies can reveal complexes of morphologically indistinguishable, but genetically very different, cryptic species, which are common among nematodes [12,13,14]. Thirdly, the patterning of population genetic structure of parasitic nematode species can inform on parasite phylogeography (see Table 1), and in so doing also clarify aspects of host phylogeography [15,16,17,18,19]. Finally, by studying the population genetics of parasitic nematodes in wild animals, ecological drivers of population genetic patterns in parasitic nematodes which may not be apparent in human- and livestock-infecting species can be identified.
The population genetics of metazoan parasites, including nematodes, has been reviewed extensively [9,10,11, 20,21,22,23,24,25], but much of this information is based on species that infect people and livestock. It is not clear how applicable these findings are to nematodes whose hosts establish their populations naturally. Parasites’ life histories, their hosts’ life histories, and the extra-host environment will all contribute to parasites’ population genetic structure [10, 11, 22, 23, 25], but little is known about the relative importance of these factors in natural ecosystems, and how they interact.
Here, we provide a resource that collates what is known about the population genetics of parasitic nematodes in natural ecosystems, that we envisage will be useful to researchers investigating the important, but little-understood, roles that parasitic nematodes play in such ecosystems. To this end, we comprehensively review the population genetics of parasitic nematodes in wild animals, including every relevant study of which we are aware. We take wild animals to be ones that establish populations without direct human management, even if they live commensally in human settlements (such as cities), or if their habitat is undergoing anthropogenic changes, since these host populations may nonetheless influence the population genetics of their parasitic nematodes in ways that hosts whose populations are managed by humans cannot. We consider the parasitic nematodes of terrestrial vertebrates, marine vertebrates, and arthropods, and draw together the evidence to present a synthesis on the factors determining the population genetics of parasitic nematodes in wild animals. We examine the population genetics of parasitic nematodes in wild animals that have undergone recent habitat change, asking how parasite populations respond to anthropogenic influences on natural systems. Finally, we assess future prospects in the study of parasitic nematode population genetics, discussing opportunities provided by high-throughput DNA-sequencing-based methods, and highlighting the importance of including extra-host stages in population genetics studies.
Parasitic nematodes of wild terrestrial vertebrates
Parasitic nematodes of marsupials
Commonly, population genetic studies of the parasitic nematodes that infect macropod marsupials reveal that what was previously classified as a single nematode species with a broad host range is actually a complex of cryptic species, each with a narrow host range (Table 2). This highlights the ability of population genetic techniques to detect species complexes. In some cases, these studies also show whether, and if so, how, populations within these cryptic species are geographically structured. For example, geographical structuring of genetic diversity was not seen in populations of Globocephaloides trifidospicularis , nor in several species within the Hypodontus macropi complex , nor in Rugopharynx australis from Macropus robustus and M. rufus , nor in several Capillaria species , but was observed in populations of H. macropi from subspecies of Macropus robustus, H. macropi from subspecies of Macropus rufogriseus  and Labiosimplex australis from M. rufogriseus . In some cases, studies have failed to detect any genetic variation within parasitic nematode species at all, such as in H. macropi from Petrogale persephone , Globocephaloides affinis from Macropus dorsalis , and several other Cloacina species , but this may be due to the very low numbers of hosts and parasites studied (Table 2).
Why do some of these parasite species have genetic population structuring while others do not? There was genetic differentiation between L. australis collected from Tasmania and Kangaroo Island, and between both of these populations and mainland Australia . Similarly, Tasmanian populations of H. macropi in M. rufogriseus showed genetic differentiation from that on mainland Australia . This suggests that marsupial hosts are ineffective at mediating nematode transmission across open water. However, a cryptic species of R. australis in M. giganteus and M. fuliginosus did not show population differentiation despite being sampled from both Kangaroo Island and mainland Australia , nor did G. trifidospicularis sampled from multiple macropod species . This may indicate ongoing gene flow in these species between Kangaroo Island and the mainland, or alternatively, that one or both of these species only recently arrived on Kangaroo Island, and the island parasite populations have yet to differentiate detectably from their mainland counterparts.
Genetic differentiation is rarely detected within marsupial-infecting nematode species collected from only mainland Australia; it is apparently absent in three cryptic species of H. macropi in Macropus dorsalis, M. agilis and Wallabia bicolor, respectively , as well as in a single species within R. australis collected from M. robustus and M. rufus . However, Queensland populations of a cryptic species of H. macropi recovered from M. robustus were found to be differentiated from those in South Australia . This may simply reflect the size of the study area; sampling locations of H. macropi in M. robustus were over 1500 km apart, compared with up to 800 km in other species. Naturally, as distances between populations increase, host-mediated transmission between the populations becomes rarer, though the exact distances over which host-mediated transmission becomes inefficient will likely vary with the vagility of host species. It should be noted that in many cases these studies involved low numbers of hosts and parasites (Table 2), and so population genetic insights gleaned from them should be treated with caution. Further work using much larger sample sizes is needed to confirm the patterns of population genetic structure in the parasitic nematodes of marsupials that many existing studies suggest.
Parasitic nematodes of terrestrial carnivores: Trichinella
Trichinella spp. have broad geographical ranges and broad host species ranges, with each species parasitizing a variety of carnivorous vertebrates. Trichinella spp. share certain population genetic characteristics, such as differentiation among infrapopulations [32, 33] (see Table 1) and low intra-specific genetic diversity [32,33,34,35,36,37,38,39]. Trichinella spp. tend to show population genetic structuring among continents; for example, genetic differentiation was found among populations of T. pseudospiralis from Australia, North America, Europe and Asia [38, 40], and among T. spiralis populations from Asia and Europe . Trichinella nelsoni from Kenya and Tanzania was differentiated from that in South Africa , but this parasite was recovered from only one host individual in each country, meaning that infrapopulation differentiation was not accounted for.
The apparent lack of population genetic differentiation in Trichinella spp. within continents matches observations made of other nematodes that parasitise large carnivorous mammals [41,42,–43], and likely arises from long-distance dispersal of hosts, which promotes high parasite gene flow. For Trichinella spp., gene flow may also be promoted by smaller hosts (such as rats and foxes), facilitating parasite gene flow among otherwise discontiguous populations of very mobile hosts.
But what factors drive the among-host differentiation and low intra-specific diversity seen in Trichinella spp.? Trichinella transmission stages remain in the muscle of their parents’ host and infection of a new host occurs by predation . This life-cycle may lead to clumped transmission of siblings, potentially resulting in differentiation among infrapopulations and promoting inbreeding. Inbreeding tends to reduce effective population size (Ne, see Table 1), leading to stronger genetic drift (see Table 1), meaning that alleles are more readily lost from the population, reducing genetic diversity. Indeed, clumped transmission of related parasites has previously been suggested to promote low genetic variation and genetic differentiation among hosts in diverse parasite taxa (reviewed in [9, 10, 21]).
In summary, the population genetics of Trichinella spp. appears to be driven by (i) highly mobile hosts and a broad host range, promoting gene flow, and (ii) its life-cycle, promoting clumped transmission of sibling parasites and so lowering Ne.
Parasitic nematodes of rodents
The limited dispersal of wild rodent individuals [45,46,47,48] might be expected to limit gene flow in their nematode parasites, resulting in genetic structure over small geographical scales. Accordingly, populations of Heligmosomoides polygyrus, a parasite of the European woodmouse (Apodemus sylvaticus), show extensive population genetic structure across the host species’ range [15, 49,50,51], with H. polygyrus populations being more strongly differentiated than those of A. sylvaticus according to mitochondrial sequence analysis of both species . Heligmosomoides polygyrus has a faster mitochondrial mutation rate and generation time than its host [15, 52], likely meaning that mitochondrial genetic drift is faster in H. polygyrus compared with A. sylvaticus and so contributing to the comparatively stronger population genetic structure of H. polygyrus.
Trichuris muris infects rats and mice (including A. sylvaticus), while T. arvicolae infects arvicoline rodents (lemmings and voles), and both Trichuris spp. are found throughout Europe. Like H. polygyrus, T. muris and T. arvicolae both show extensive population genetic structure across their geographical range, as determined by analysis of both mitochondrial and nuclear loci [53,54,55]. Indeed, broadly similar patterns of population genetic structure were observed in H. polygyrus and both Trichuris spp., with a delineation between eastern and western populations, and greater diversity in southern populations, compared with northern. These patterns may reflect range expansion of the rodent hosts from southern refugia during the last ice age, at least 12,000 years ago [15, 53,54,55]. Stronger population genetic structuring and a smaller geographical range was observed in H. polygyrus than in Trichuris spp. [15, 53,54,55]. This may be partly due to faster genetic drift in H. polygyrus than in Trichuris spp., arising from a shorter generation time in the former compared with the latter (~14 days and 50–60 days, respectively) [56, 57], or may reflect the broader host range of Trichuris spp. A broader host range may allow a parasite to occupy and traverse a wider range of environments, potentially increasing gene flow rates and slowing genetic drift .
The population genetics of Angiostrongylus cantonensis and A. malaysiensis, parasites of rodents of the family Muridae , has been extensively studied, with genetic structure detected on both small and large geographical scales [59,60,61,62,63,64,65,66,67,68,69]. However, interpretation of these findings is made difficult by the recent discovery of at least two sympatric cryptic species within A. cantonensis, one of which may be conspecific with individuals identified as A. malaysiensis [70, 71]. It is now not clear whether the population genetic structure detected previously really represents the geographical distribution of genetic variants in a single species, or rather results from the accidental sampling of multiple reproductively isolated species. Nevertheless, if this cryptic speciation is taken into account, the population genetic data of A. cantonensis (sensu stricto) and A. malaysiensis can be examined. In both species, population genetic structure was detected among provinces in Thailand . It is likely that limited vagility in both the definitive rodent host and the intermediate snail host limits gene flow in Angiostrongylus spp. over large distances.
Other studies have investigated parasitic nematode population genetic structure at finer geographical scales. For example, Neoheligmonella granjoni, a parasite of multimammate mice (Mastomys spp.), was sampled from M. natalensis and M. erythroleucus within a 70 km2 rural area of Senegal, and genotyped at 10 microsatellite loci . These data revealed an absence of genetic population structure, with alleles being distributed homogenously among sampling sites. This may be due to the relatively high dispersal of M. erythroleucus, which will promote gene flow across the study area, including among populations of the much more sedentary host, M. natalensis [72, 73]. A lack of population genetic structure over fine geographical scales (e.g. within a 100 km2 area) was also seen in Longistriata caudabullata, a nematode that parasitises Blarina sp. shrews , showing that while parasitic nematodes of rodents and other small mammals show strong genetic structure at broad geographical scales, at finer scales, gene flow can be sufficient to genetically homogenise populations.
Strongyloides ratti is a parasite of brown rats, Rattus norvegicus, and shows little genetic differentiation among UK sampling sites ~20–250 km apart . This may indicate that R. norvegicus dispersal is sufficient to genetically homogenise the S. ratti population at these scales. While S. ratti did not show genetic differentiation among host populations, there was differentiation among infrapopulations . Strongyloides spp. are unusual because the parasitic adults reproduce clonally, so that all of a single parasite’s offspring are genetically identical , and along with clumped transmission of clonal siblings , this may lead to the observed among-host differentiation.
Analysis of Syphacia stroma and H. polygyrus from the same A. sylvaticus host individuals showed that S. stroma has substantially lower genetic diversity and higher population differentiation than H. polygyrus . Syphacia stroma has haplodiploid sex determination, in which haploid males develop from unfertilised eggs produced by diploid females, while in H. polygyrus males and females are both produced sexually. Mating system is recognised as an important factor in parasite population genetics [10, 11, 23], and so the different mating systems of H. polygyrus and S. stroma may explain their different population genetic structures. Haplodiploidy lowers Ne by reducing the number of individuals contributing to the next generation (because males are produced from the mother’s genetic material only), and this may lead to greater genetic drift in S. stroma compared with H. polygyrus. Syphacia stroma and H. polygyrus have broadly similar generation times  and share a host, so their different mating systems emerge as likely important factors behind their different population genetics. Aspects of life history such as mating system have not been extensively studied in parasitic nematodes of wild animals, and further studies in this area may contribute to our understanding of population genetics in other parasitic nematode species.
In summary, the population genetics of parasitic nematodes in wild rodents appears to be defined largely by hosts’ low dispersal ranges. However, different patterns of population genetic structure among parasite species sharing a host species suggest that parasite mating system and generation time are also influential.
Parasitic nematodes of ungulates
Ungulate (hoofed mammal) individuals travel over much greater distances than rodents, and so may facilitate comparatively greater gene flow of their parasitic nematodes. Ostertagia gruehneri and Marshallagia marshalli, both parasites of reindeer (Rangifer tarandus), show a lack of population genetic structuring [80, 81], a pattern similar to that of Teladorsagia boreoarcticus in muskoxen (Ovibos moschatus) . In contrast, Mazamastrongylus odocoilei, a parasite of white-tailed deer (Odocoileus virginianus), showed genetically structured populations . This difference may reflect the more rapid evolution of mtDNA, used to study M. odocoilei, compared with the internal transcribed spacer (ITS, see Table 1) sequences used for O. gruehneri and M. marshalli. However, species-specific differences in host ecology may also contribute to the different patterns of population genetic structure seen among O. gruehneri, M. marshalli and M. odocoilei - reindeer have large home ranges and are partially migratory , and so may provide more opportunities for gene flow in their parasitic nematodes compared with the more sedentary white-tailed deer .
Dictyocaulus eckerti is a parasite of several species of deer (Cervus spp. and Dama spp.). Analysis of mitochondrial sequence data found weak genetic structuring in D. eckerti , while D. capreolus (specific to roe deer, Capreolus capreolus), had comparatively lower genetic diversity and more strongly genetically structured populations when sampled sympatrically . Dictyocaulus capreolus is susceptible to population bottlenecks if roe deer numbers fall, whereas D. eckerti can weather a crash in the population of any one host species by persisting in other host species, and thereby maintain a high census size (N, see Table 1). High genetic diversity and a genetically unstructured populations is also observed in Trichostrongylus axei, which parasitises diverse wild ungulate species , suggesting an association between these population genetic traits and broad host range. Differences in host behaviour may also contribute to the differences in the population genetics of D. eckerti and D. capreolus; specifically, D. eckerti may have higher gene flow than D. capreolus because of the territorial nature of roe deer, which limits the geographical distances they cover.
Parasitic nematodes of reptiles and amphibians
Spauligodon anolis infects anole lizards (Anolis spp.), while Parapharyngodon cubensis is a species complex (P. cubensis A, P. cubensis B and P. cubensis C) together infecting a broad range of lizards and snakes. Study of the population genetics of these nematodes, sampled from various Caribbean Anolis spp. hosts, found that genetic diversity was partitioned both among and within islands . However, S. anolis populations were more strongly genetically differentiated than populations of P. cubensis A or P. cubensis B, likely because S. anolis has a narrow host species range made up of poor dispersers , while the species of the P. cubensis complex each make use of a wider range of hosts, among which may be more mobile host species . In contrast, cryptic species within Spauligodon atlanticus, parasites of Gallotia spp. lizards, all showed strong genetic structuring within and among islands of the Canary Isles, despite differing in the extent of their host range . This may be because the geographical ranges of the host species of S. atlanticus do not overlap, precluding nematode gene flow between them.
Population genetic analysis of Rhabdias ranae, a parasite of the northern leopard frog (Lithobates pipiens), revealed low microsatellite heterozygosity, differentiation among infrapopulations and population genetic structure at a very fine scale, with differentiation emerging among ponds less than 1 km apart . Rhabdias ranae is a specific parasite of L. pipiens and lacks an intermediate host, so its dispersal is likely mediated almost entirely through L. pipiens movement. Hence, sibling extra-host stages are likely to remain clumped in the environment and infect a new host together, explaining infrapopulation differentiation. If L. pipiens habitually visit the same locations (e.g. show fidelity to a particular breeding pond), then they may even be re-infected with the offspring of their own parasites . Such a habit might also explain differentiation among ponds, if the same cohort of frogs routinely utilise a particular pond . Low heterozygosity is likely a product of inbreeding, arising from the life-cycle of R. ranae, which includes a self-fertilising hermaphroditic stage.
Parasitic nematodes of terrestrial birds: Trichostrongylus tenuis
Trichostrongylus tenuis is a strongylid parasite of galliform and anseriform birds, and is particularly prevalent in red grouse (Lagopus lagopus scotica). Population genetic analysis of T. tenuis in UK red grouse revealed high microsatellite diversity, and a lack of population genetic structure among host individuals and among geographically separated populations [92, 93]. This lack of population genetic structure is likely due to the very high prevalence and infection intensity of this parasite , presumably leading to a high Ne, so rendering genetic drift very slow. Population genetics can also be used to study parasite dispersal. For example, a lack of genetic differentiation between T. tenuis in a goose in Iceland and those in UK grouse suggests long distance T. tenuis gene flow. Some waterfowl species, such as the pink-footed goose (Anser brachyrhynchus), migrate between the UK and Iceland , presenting a possible avenue for T. tenuis gene flow between these countries.
Parasitic nematodes of aquatic vertebrates
Parasitic nematodes of marine mammals and birds
Most nematode parasites of marine mammals and birds are trophically transmitted among intermediate hosts before reaching the definitive host . As hosts of each trophic level are likely to consume multiple infected hosts in the lower trophic levels, hosts will accumulate parasites from a variety of sources, and so definitive host individuals will probably sample widely from the parasite population. This may lead to genetically diverse parasite infrapopulations that obviate inbreeding and promote high Ne . Because many marine fish, mammal and bird hosts travel large distances [97, 98], gene flow of their parasitic nematode populations is expected to be high, suggesting that these nematode populations will show little genetic structuring.
Many parasitic nematodes infecting marine vertebrates do indeed show little population genetic structure. Anisakis simplex is a complex of several cryptic species with different geographical and definitive host ranges [99,100,101]. Population genetic structure has rarely been observed within species of the A. simplex complex (Table 3), and it is likely that earlier reports of extensive genetic structure in A. simplex  resulted from inadvertent sampling of multiple species. A similar lack of population genetic structure has been observed in a variety of other nematodes with similar life histories, including other Anisakis spp. in pinnipeds and cetaceans, Contracaecum spp. from a variety of birds and mammals, and Pseudoterranova spp. from pinnipeds (Table 3). However, Anisakis simplex C may be an exception, with one study detecting genetic differentiation between northern and southern hemisphere populations . Intra-taxon genetic diversity of parasitic nematodes of marine mammals in the southern hemisphere is generally greater than in the northern hemisphere, perhaps due to comparatively lower habitat disturbance (e.g. fishing, pollution) in the southern hemisphere .
Uncinaria sanguinis, a parasite of the Australian sea lion (Neophoca cinerea), requires adult female hosts on breeding beaches to complete its life-cycle , and female hosts always return to the beach they were born on to breed . One might therefore expect U. sanguinis to show genetic differentiation among host breeding beaches, but in fact no population genetic structure was observed in U. sanguinis at all , and a similar situation is seen in Uncinaria lucasi infecting northern fur seals (Callorhinus ursinus) . This may indicate that the life-cycle of Uncinaria spp. is not fully understood and that transmission also occurs in other ways - male sea lions do move among breeding beaches , so transmission involving males could homogenise parasite population genetic structure. Hence, population genetic studies can suggest hypotheses about transmission cycles of parasitic nematodes that might otherwise be unexpected.
Parasitic nematodes of fish
Some fish species travel around the globe, while others spend their whole lives in a small home patch, and this diversity in movement behaviour is likely to affect the population genetics of their parasitic nematodes. Hysterothylacium aduncum is a poorly-defined nematode species complex that infects a broad range of marine fish species . ITS sequence data of H. aduncum from sprats (Sprattus sprattus) in western Europe showed two genetically distinct populations: one in the English Channel and the Bay of Biscay, and one in the Mediterranean and North Sea . The geographical separation of the Mediterranean and North Sea (and that they are separated by the English Channel and Bay of Biscay), makes this parasite population genetic structure peculiar, and it contrasts with the population genetics of the sprats . Potentially, another host species may be responsible for genetically homogenising the H. aduncum populations in the Mediterranean and North Sea via migration, and sampling of H. aduncum from other hosts is needed to test this hypothesis. In contrast with H. aduncum, there was no genetic structure in Hysterothylacium fabri within the Mediterranean, when considering either geography or host fish species .
Parasites of fish in discontiguous water bodies can become genetically distinct. For example, splitfin fishes (several genera within the Goodeidae) live in a series of unconnected lakes in Mexico, and their parasite, Rhabdochona lichtenfelsi, shows strong genetic differentiation among lakes, with the degree of differentiation correlating with the time since the lakes became separated . In contrast, populations of the yellowhead catfish (Pelteobagrus fulvidraco) parasite Procamallanus fulvidraconis in isolated lakes were not significantly genetically different from each other . These lakes were connected until the 1950s , so there may have been insufficient time for the parasite populations to diverge genetically. Camallanus cotti parasitizes a variety of freshwater fish species, and it showed no population genetic structure among the Yangtze and Minjiang river systems (geographically close and possibly occasionally connected by flood water), though populations from the Pearl River were distinct .
Collectively, studies of the population genetics of parasitic nematodes in aquatic environments reveal that their population genetic structures emerge at the scale over which hosts move, with genetically unstructured populations being common. Population genetic structure can emerge in these parasites when populations are restricted to isolated water bodies, or where host movement is constrained.
Nematode parasites of arthropods
Virtually all invertebrate taxa are infected by parasitic nematodes, but the life histories of these nematodes are often poorly understood, and their population genetics barely explored. The insect parasite Heterorhabditis marelatus has a low level of mitochondrial genetic diversity and shows extreme population genetic structuring among sample sites (7 to 890 km apart) . This may arise from very low gene flow in H. marelatus, since the extreme pathogenicity of H. marelatus kills hosts before they can carry their parasites far, preventing host movement from contributing significantly to parasite dispersal . The life-cycle of H. marelatus may also contribute to its strong population genetic structure; Heterorhabditis spp. infections are initiated by juveniles which, upon maturation into hermaphrodites, self-fertilise to produce males and further hermaphrodites that continue to reproduce on the host’s cadaver . This life-cycle promotes frequent founder effects (when an infective juvenile invades a host) and inbreeding (self-fertilisation and sib-mating), together driving low Ne. Low genetic diversity and population genetic structure was also observed in Strelkovimermis spiculatus, sampled from mosquito larvae (Aedes spp. and Culex pipiens), with genetic differentiation observed among ponds ~7 to 14 km apart . In contrast, no population genetic structure was observed in Isomermis lairdi, a parasite of larval blackflies (Simulium spp.), when sampled from three rivers and multiple host species . That the rivers were connected likely facilitates gene flow of I. lairdi, resulting in less structured populations compared with S. spiculatus.
Thaumamermis zealandica, a parasite of the sandhopper (Bellorchestia quoyana, a beach-dwelling amphipod), showed a complete absence of genetic diversity in three mitochondrial protein-coding genes when sampled from numerous hosts along an ~580 km stretch of New Zealand coast . This could result from (i) panmixia among sampling sites and an extremely low Ne in the entire population, such that genetic drift affects T. zealandica at all sampling sites as a single population; (ii) a very recent population bottleneck; (iii) an extremely low mitochondrial mutation rate; or (iv) a combination of these factors . Extremely low genetic variation in mitochondrial protein-coding genes was also seen in the woodlouse (Armadillidium vulgare) parasite Thaumamermis cosgrovei , suggesting that a low mutation rate in mitochondrial protein-coding genes may be a common feature of Thaumamermis spp.
RAPD (see Table 1) analysis of Blatticola blattae, a parasite of cockroaches (Blattella germanica), showed that the parasite population genetic structure closely mirrored that of the host, with both showing differentiation among buildings within cities, and among cities 900 km apart . This strong genetic structuring likely reflects limited dispersal in cockroaches, promoting low gene flow in both parasite and host. Unlike H. marelatus, B. blattae is not markedly pathogenic and individuals form long-term associations with their hosts , so that there is time for host movement to mediate parasite gene flow.
Among parasitic nematodes of arthropods, then, pathogenicity to the host may influence the parasites’ population genetic structure, as parasites that kill their hosts very quickly cannot rely on host movement for dispersal and gene flow. However, where the arthropod host has a very small home range and does not disperse far, even largely non-pathogenic parasites may have strongly structured populations, as seen in B. blattae . Nevertheless, our knowledge of the population genetics of parasitic nematodes of arthropods is very incomplete, and future work analysing a broader range of both host and parasite life histories is needed if we are to better understand the factors influencing their population genetics.
Influence of anthropogenic disruption on parasitic nematode population genetics
Human activities have affected the geographical ranges of many host species, either shrinking a range through habitat loss, or increasing it through introduction of individuals into new regions. In many cases, the timing and extent of range changes are well documented, offering an opportunity to study how changes in host population size and connectivity shape parasitic nematode population genetics.
Baylisascaris schroederi is a nematode thought to be specific to giant pandas (Ailuropoda melanoleuca). Sequence analysis of both ITS and mtDNA have shown a lack of genetic differentiation among B. schroederi from geographically isolated panda populations [124,125,126,127], which is surprising because pandas do not migrate among populations, and pandas from each of these populations are themselves genetically differentiated . The likely explanation for this difference between host and parasite population genetics is that B. schroederi has a much larger Ne than its host  and so undergoes population differentiation more slowly. Thus, while there may have been time for panda populations to differentiate through genetic drift in the 200 years since habitat fragmentation began , drift may not have been fast enough to yet differentiate B. schroederi populations. It has also been suggested that B. schroederi gene flow may occur among panda populations in the absence of panda movement, for example through association with a presently unknown paratenic host .
An analogous situation is seen in Trypanoxyuris minutus and T. atelis, parasites of the primates Alouatta spp. and Ateles geoffroyi, respectively, in Mexico. Since 1940, on-going forest fragmentation has created discontiguous host populations, among which host migration is very rare . Despite this, mitochondrial sequence analysis of Trypanoxyuris spp. showed that parasite populations in different forest fragments were not genetically differentiated . There are two non-mutually-exclusive explanations for this; unexpectedly high Trypanoxyuris gene flow among forest fragments, and/or the failure of Trypanoxyuris populations to detectably differentiate since becoming reproductively isolated. The latter explanation assumes that Trypanoxyuris populations were genetically unstructured prior to forest fragmentation, and this seems plausible as parasitic nematodes from a range of wild primates show limited population genetic structure when looking within host species [133,134,135,136,137,138,139].
The population genetics of Baylisascaris procyonis infecting raccoons (Procyon lotor) has been studied using both ribosomal and microsatellite loci in its native range [140, 141], with microsatellite loci revealing genetic differentiation across the Grand River (Michigan, USA). Among invasive B. procyonis populations in Germany, two well-differentiated clades have been detected by both ITS and mitochondrial sequence analysis, suggesting two independent introductions of B. procyonis into Germany . Both German B. procyonis clades showed very low genetic diversity, likely the result of population bottlenecks (see Table 1) in the founding populations . Low genetic diversity was seen also in Rhabdias pseudosphaerocephala in invasive cane toads (Bufo marinus) in Australia , and in Passalurus ambiguus in invasive rabbits (Oryctolagus cuniculus) in China , and is typical due to founder effects in these introduced species .
The nematode Spirocamallanus istiblenni was introduced to the Hawaiian archipelago with one of its hosts, the bluestripe snapper (Lutjanus kasmira). Population genetic analysis of S. istiblenni confirmed that the parasite originated from French Polynesia and showed that the introduced population was less genetically diverse than the native population . Population genetic data also provided evidence that the introduced S. istiblenni has transmitted to native fish, shown by a lack of genetic differentiation between parasites from introduced and native hosts. Population genetic investigations into Camallanus cotti, invasive in Hawaiian stream fishes, revealed its probable invasion history, suggesting an initial introduction in O’ahu, where genetic diversity was highest, and subsequent migration to other islands in the archipelago . Comparisons of the data in this study with data from C. cotti in its native range  showed that genetic diversity in introduced C. cotti was reduced compared with native populations, once again demonstrating the effects of population bottlenecks in introduced parasitic nematode populations.
Anguillicoloides crassus and its host, the Japanese eel (Anguilla japonica), were introduced from Asia to North America and Europe, and since then A. crassus has spread rapidly in European and American eels (Anguilla anguilla and A. rostrata, respectively), causing severe pathology in these naïve host species . Population genetic studies have revealed multiple, distinct lineages of invasive A. crassus, suggesting multiple introduction events from different source populations [149, 150]. Furthermore, a southern to northern clinal decrease in its genetic diversity is seen in Europe, suggesting that A. crassus was introduced in southern Europe and has since spread northwards . Hence, the population genetics of A. crassus has revealed its introduction history. Infrapopulation differentiation in A. crassus was studied in two European rivers, one of which was regularly artificially restocked with eels from a variety of sources and one in which eels had arrived by natural dispersal . This showed that in the restocked river, A. crassus had high genetic diversity among hosts and a substantial deviation from Hardy-Weinberg equilibrium (see Table 1), while in the river with natural recruitment, there was no among-host structuring or deviation from Hardy-Weinberg equilibrium. These contrasting patterns were thought to be because the introduced eels had retained A. crassus infrapopulations reflective of their genetically distinct source populations, while A. crassus in the river with natural recruitment are derived from a single population that was already at Hardy-Weinberg equilibrium .
As environments continue to change, the ranges of parasitic nematodes will change too. The population genetics of parasitic nematodes currently undergoing such range changes show that (i) invasive parasitic nematodes are likely to have low genetic diversity (due to population bottlenecks); and (ii) that host populations are likely to lose diversity more rapidly following habitat fragmentation than parasitic nematode populations (due to the comparatively smaller Ne of hosts). This latter effect may have a consequence for parasite ecology, since hosts will differ in their genetic resistance to parasites, and genetic bottlenecks of host populations may therefore lead to altered degrees of parasitism. Supporting this, Trypanoxyuris spp. and B. schroederi all show genetic evidence of recent population expansion despite marked declines in host population size [127, 132], suggesting that prevalence and/or intensity of infection has increased since habitat fragmentation. Hence, population genetic analysis can inform on the biology of parasitic nematodes undergoing changes in range, and can be used to make predictions about how parasite populations might respond genetically to future range changes.
Prospects in nematode population genetics
Each of the methods routinely used to analyse parasitic nematode population genetics detects variation in a small number of loci , and it is often not clear how representative these loci are of a genome more widely. High-throughput sequencing techniques can be employed to interrogate large portions of the genome, thus reducing the effect of bias at any one genomic region. Other advantages include the ability to screen the genome for regions under selection, and the chance to analyse population genetic structure at several scales simultaneously. For example, highly polymorphic (see Table 1) genomic regions can inform on structure at very local scales, while more conserved regions will be appropriate for studying the relationships of more divergent populations. Genome-wide sequencing has been used to assess population genetics in non-nematode helminths, allowing detailed insight into parasite population genetic structure , and there is no reason why such insights should not be possible in parasitic nematodes. These techniques usually require a reference genome, which often will not be available a priori for non-model parasitic nematodes infecting wild animals. However, rapid advances in the sequencing and assembly of nematode genomes mean that it may often be feasible to generate a reference genome for the species in question .
In restriction-associated DNA sequencing (RADSeq) the genome of an individual is digested with a restriction endonuclease, and the resulting fragments are size-filtered and then sequenced . Random distribution of restriction endonuclease sites across the genome ensures that the sequenced fragments are representative of the whole genome. Double-digest RADSeq (ddRADSeq) is a related technique that uses two restriction endonucleases . While (dd)RADSeq data can be used without a reference genome assembly , having a reference means that non-target DNA can be detected and excluded. (dd)RADSeq approaches have not yet been used in the population genetic analysis of parasitic nematodes, but have been used in several other animal species [9, 157] including the free-living nematode Caenorhabditis elegans . The study in C. elegans not only revealed the population genetics of this species, but also its recent evolutionary history, finding that the low genetic diversity of C. elegans likely arose from recent selective sweeps (see Table 1), in which a few beneficial alleles drove large swathes of the genome to near-fixation (see Table 1) due to extensive linkage disequilibrium (see Table 1) . If (dd)RADSeq were used to study the population genetics of parasitic nematodes, we could expect similar insights into the biology and evolutionary history of these species.
In whole-genome sequencing, individuals are genotyped at virtually every locus polymorphic among samples. This allows the relationships among individuals, and hence the genetic structure of populations, to be resolved at the finest possible scale , and gives more power to make inferences about the evolutionary processes acting on a species [160, 161]. There are currently obstacles to routinely generating whole-genome sequence data. Firstly, sequencing the genomes of multiple individuals is often still expensive, such that there is a trade-off between the number of individuals sequenced and the genome coverage of each individual. However, population genetic techniques are robust to low genome coverage, with coverage of as little as two-fold having been applied . Secondly, the small physical size of some nematodes can make it difficult to extract sufficient DNA for sequencing. Whole-genome amplification (WGA) prior to sequencing can ameliorate this difficulty , though not without some bias in genome coverage .
However, the difficulties of whole-genome sequencing are worth overcoming, due to the wealth of information it can provide. Of particular importance is the ability of whole-genome sequencing to identify genes that are under selection, and in the case of parasites, these genes may be relevant to pathogenicity, epidemiology and control. For example, whole-genome sequencing of the malaria parasite Plasmodium falciparum has found that genes that function in evasion of host immunity and in resistance to drugs show signs of selection . Similar studies in parasitic nematodes are warranted, as anthelmintic resistance is an increasingly serious problem in both agricultural and medical settings, and the biochemical mechanisms of resistance are often poorly understood [20, 166]. Genes that show strong signatures of directional selection in anthelminthic-exposed populations are good candidates for genes conferring anthelmintic resistance .
Whole-genome sequencing has already been used to study the population genetics of parasitic nematodes that infect people, demonstrating among-host differentiation in Wuchereria bancrofti , and detecting differentiation in Strongyloides stercoralis among host individuals and populations . However, these studies used small numbers of hosts and are limited in scope. Hence, there is now a need for studies on a comparatively larger scale that examine the distribution of polymorphisms within genomes as well as among them. In an intriguing recent study, whole-genome sequencing was used to interrogate the population genetics of the plant-parasitic nematode Heterodera glycines without the use of a reference genome . This study, which made use of the UNEAK bioinformatics pipeline , was able to not only elucidate the population genetic structure of H. glycines, but also to identify genetic variants showing signatures of selection. Whether a reference assembly is used or not, whole-genome sequencing studies will improve our understanding of how parasitic nematodes respond to natural selection pressures. As the climate changes, parasitic nematodes will encounter novel selection pressures, and how they respond to these pressures may have important consequences, not only for host species but for the ecosystem as a whole.
Environmental DNA (eDNA) analysis
Parasitic nematodes typically have extra-host transmission stages in their life-cycle, and as these stages are the pool from which the next generation of parasites will arise, they contribute directly to Ne. Differences in the number, spatial distribution and temporal distribution of these stages may therefore influence the rate of genetic drift in a population, and hence the population genetic structure more widely. Despite their likely importance in both population genetics and epidemiology, very little is known about the ecology of extra-host stages, and what is known largely pertains to human or livestock pathogens [172, 173]. This knowledge gap is principally due to the difficulty in sampling and identifying extra-host nematode stages in the environment. Sequencing of environmental DNA (eDNA, see Table 1) may be a solution to this problem [174, 175]. DNA-based identification of parasitic nematodes stages in host faeces - in essence a form of eDNA - is often used to diagnose infection in livestock . Adaptation of these techniques for the detection of parasitic nematode stages in soil and water (where target DNA concentrations will often be lower) would incorporate extra-host transmission stages into analysis of population genetics of parasitic nematodes infecting wild animals.
Synthesis and outstanding research questions
As with all species, the population genetics of parasitic nematodes in wild animals is ultimately determined by the (i) rate of gene flow among populations and (ii) strength of genetic drift within populations. Nematodes have very limited dispersal ability of their own, so that they are largely dependent on their hosts for dispersal. Therefore, in principle, nematode populations should ultimately come to be structured at the scale over which hosts move. Broadly speaking, this rule is largely followed - there is very limited population genetic structuring of parasitic nematodes infecting highly mobile hosts, such as ocean-going mammals , while the parasites of small, sedentary rodents, for example, generally have highly structured populations .
Nematode species that use more than one host species often have less structured populations than the movement of any one host species would predict. This has been observed in Trichinella spp., which shows gene flow at the continental scale , in Dictyocaulus eckerti, a generalist parasite of deer , and in Neoheligmonella granjoni [72, 73]. The use of multiple mobile host species allows nematodes to traverse or colonise a wider range of habitats than would be possible using only a single host, and this tends to promote parasite gene flow. In contrast, highly pathogenic parasitic nematodes, such as Heterorhabditis marelatus, may show reduced gene flow if they significantly hamper their host’s movement .
In the absence of gene flow all populations will ultimately diverge genetically due to genetic drift, but this process takes time. How much time depends on the strength of genetic drift, in turn dictated by (among other things) Ne, and so Ne is a major determinant in population genetic structure. Ne is rarely measured directly but can be estimated from known aspects of parasite and host life history. For example, haploid males in Syphacia stroma likely reduce Ne in comparison with the fully diploid Heligmosomoides polygyrus, with which S. stroma shares a host , while the much lower infection prevalence and intensity of Dictyocaulus viviparus compared with Ostertagia ostertagi in domestic cattle means that the former likely has lower Ne [177, 178]. In both cases, the species presumed to have the lower Ne has stronger population genetic structuring. Naturally, parasites with a fast generation time will undergo more rapid genetic drift (and so faster population divergence) than parasites with a slow generation time. Indeed, the significantly faster generation time of H. polygyrus compared with Trichuris muris may contribute to the comparatively more strongly genetically structured populations of the former when both species are analysed from the same host individuals [15, 53, 54].
Parasite life history traits such as host range, reproductive strategy, generation time, and the prevalence, intensity and pathogenicity of infection in the host are therefore all important in determining the population genetics of parasitic nematodes. These factors interact with host ecology, and in particular, host movement behaviour, to establish patterns of parasite population genetic structure. However, the relative importance of parasite life history traits remains poorly understood. Animals are commonly infected by more than one species of nematode, and this fact could be exploited to better understand population genetics in nematodes infecting wild animals. For example, comparison of life history traits among co-infecting parasites would allow their effects on parasite populations genetics to be separated from host-dependent effects.
The abundance and spatial and temporal dynamics of extra-host stage parasitic nematodes in the environment remains almost entirely unknown. There have been some attempts to study the extra-host stages of nematodes infecting domestic livestock , but the findings may not be fully applicable to species infecting wild animals. Understanding the ecology of extra-host parasite stages is important for conservation of wild hosts, for monitoring the threat of zoonotic infection, and for our understanding of ecosystem processes. Particularly mysterious is the role of extra-host stages on the population genetics of parasitic nematodes, and future work must address this to complete our understanding of the population genetics of parasitic nematodes in wildlife. eDNA applications may be the only means of sampling extra-host nematode stages with sufficient rigour to understand their contribution to population genetics.
Looking to the future, the use of high-throughput sequencing-based methods (dd)RADSeq and whole-genome sequencing) will dramatically improve the resolution and accuracy with which population structure can be detected. Furthermore, whole-genome sequencing will allow other aspects of parasitic nematode genomes, such as the size and nature of selection, and the extent of linkage disequilibrium, to be interrogated, and this knowledge will improve our understanding of parasitic nematode biology.
The population genetics of parasitic nematodes in wild animals is determined by a combination of host ecology - especially host movement behaviour - and parasite life history. Studying the population genetics of parasitic nematodes of wild animals can reveal how their populations respond to selective pressures. With this information, we can assess the risk parasitic nematodes pose to natural ecosystems and to the health of humans and domestic animals, as anthropogenic activities drive environmental changes and changes in species’ geographical ranges. Our understanding of the population genetics, biology and evolutionary history of parasitic nematodes will be improved if investigators incorporate extra-host transmission stages and take advantage of high-throughput DNA-sequencing technologies in future studies.
Double-digest restriction-associated DNA sequencing
Internal transcribed spacer
Effective population size
Polymerase chain reaction
Restriction-associated DNA sequencing
Random amplified polymorphic DNA
Dobson AP. The population biology of parasite-induced changes in host behaviour. Q Rev Biol. 1988;63:139–65.
Hurd H. Physiological and behavioural interactions between parasites and invertebrate hosts. Adv Parasit. 1990;29:271–318.
Hudson PJ, Dobson AP, Lafferty KD. Is a healthy ecosystem one that is rich in parasites? Trends Ecol Evol. 2006;21:381–5.
Frainer A, McKie BG, Amundsen P-A, Lafferty KD. Parasitism and the biodiversity-functioning relationship. Trends Ecol Evol. 2018;33:260–8.
Kirk RS. The impact of Anguillicola crassus on European eels. Fisheries Manag Ecol. 2003;10:385–94.
Zhang J-S, Daszak P, Huang H-L, Yang G-Y, Kilpatrick AM, Zhang S. Parasite threat to panda conservation. EcoHealth. 2008;5:6–9.
Sakanari JA, McKerrow JH. Anisakiasis. Clin Micriobiol Rev. 1989;2:278–84.
Sorvillo F, Ash LR, Berlin OG, Yatabe J, Degiorgio MSA. Baylisascaris procyonis: an emerging helminthic zoonosis. Emerg Infect Dis. 2002;8:355–9.
Gilabert A, Wasmuth JD. Unravelling parasitic nematode natural history using population genetics. Trends Parasitol. 2013;29:438–48.
Gorton MJ, Kasl EL, Detwiller JT, Criscione CD. Testing local-scale panmixia provides insights into the cryptic ecology, evolution, and epidemiology of metazoan animal parasites. Parasitology. 2012;139:981–97.
Criscione CD, Poulin R, Blouin MS. Molecular ecology of parasites: elucidating ecological and microevolutionary processes. Mol Ecol. 2005;14:2247–57.
Blouin MS. Molecular prospecting for cryptic species of nematodes: mitochondrial DNA versus internal transcribed spacer. Int J Parasitol. 2002;32:527–31.
Nadler SA, Pérez-Ponce de León G. Integrating molecular and morphological approaches for characterizing parasite cryptic species: implications for parasitology. Parasitology. 2011;138:1688–709.
Pérez-Ponce de León G, Nadler SA. What we don’t recognize can hurt us: a plea for awareness about cryptic species. J Parasitol. 2010;96:453–64.
Nieberding C, Morand S, Libois R, Michaux JR. A parasite reveals cryptic phylogeographic history of its host. P R Soc B. 2004;271:2559–68.
Galbreath KE, Hoberg EP. Return to Beringia: parasites reveal cryptic biogeographic history of North American pikas. Proc R Soc B Biol Sci. 2012;279:371–8.
Wickström LM, Haukisalmi V, Varis S, Hantula J, Fedorov VB, Henttonen H. Phylogeography of the circumpolar Paranoplocephala arctica species complex (Cestoda: Anoplocephalidae) parasitizing collared lemmings (Dicrostonyx spp.). Mol Ecol. 2003;12:3359–71.
Wickström LM, Hantula J, Haukisalmi V, Hentonnen H. Genetic and morphometric variation in the Holarctic helminth parasite Andrya arctica (Cestoda, Anoplocephalidae) in relation to the divergence of its lemming hosts (Dicrostonyx spp.). Zool J Lind Soc-Lond. 2001;131:443–57.
Koehler AVA, Hoberg EP, Dokuchaev NE, Tranbenkova NA, Whitman JS, Nagorsen DW, Cook JA. Phylogeography of a Holarctic nematode, Soboliphyme baturini, among mustelids: climate change, episodic colonization, and diversification in a complex host-parasite system. Biol J Linn Soc. 96:651–63.
Gilleard JS, Redman E. Genetic diversity and population structure of Haemonchus contortus. Adv Parasitol. 2016;93:31–68.
Nadler SA. Microevolution and the genetic structure of parasite populations. J Parasitol. 1995;81:395–403.
Mazé-Guilmo E, Blanchet S, McCoy KD, Loot G. Host dispersal as the driver of parasite genetic structure: a paradigm lost? Ecol Lett. 2016;19:336–47.
Vázquez-Prieto S, Vilas R, Paniagua E, Ubeira FM. Influence of life history traits on the population genetic structure of parasitic helminths: a minireview. Folia Parasitol. 2015;62:060.
Anderson TJC, Blouin MS, Beech RN. Population biology of parasitic nematodes: applications of genetic markers. Adv Parsitol. 1998;41:219–83.
Barret LG, Thrall PH, Burdon JJ, Linde CC. Life history determines genetic structure and evolutionary potential of host-parasite interactions. Trends Ecol Evol. 2008;23:678–85.
Fazenda IP, Beveridge I, Chilton NB, Jex AR, Pangasa A, Campbell BE, et al. Analysis of genetic variation in Globocephaloides populations from macropodid marsupials using a mutation scanning-based approach. Electrophoresis. 2009;30:2758–64.
Chilton NB, Beveridge I, Andrews RH. Detection by allozyme electrophoresis of cryptic species of Hypodontus macropi (Nematoda: Strongyloidea) from macropodid marsupials. Int J Parasitol. 1992;22:271–9.
Chilton NB, Andrews RH, Beveridge I. Genetic evidence for a complex of species within Rugopharynx australis (Mönnig, 1926) (Nematoda: Strongyloidea) from macropodid marsupials. Syst Parasitol. 1996;34:125–33.
Zhu X, Spratt DM, Beveridge I, Haycock P, Gasser RB. Mitochondrial DNA polymorphism within and among species of Capillaria sensu lato from Australian marsupials and rodents. Int J Parasitol. 2000;30:933–8.
Chilton NB, Huby-Chilton F, Smales LR, Gasser RB, Beveridge I. Genetic divergence between island and continental populations of the parasitic nematode Labiosimplex australis in Australia. Parasitol Res. 2009;104:229–36.
Chilton NB, Huby-Chilton F, Johnson PM, Beveridge I, Gasser RM. Genetic variation within species of the nematode genus Cloacina (Strongyloidea: Cloacininae) parasitic in the stomachs of rock wallabies, Petrogale spp. (Marsupialia: Macropodidae) in Queensland. Aust J Zool. 2009;57(1):10.
Dunams-Morel DB, Reichard MV, Toretti L, Zarlenga DS, Rosenthal BM. Discernible but limited introgression has occurred where Trichinella nativa and the T6 genotype occur in sympatry. Infect Genet Evol. 2012;12:530–8.
La Rosa G, Marucci G, Rosenthal BM, Pozio E. Development of a single larva microsatellite analysis to investigate the population structure of Trichinella spiralis. Infect Genet Evol. 2012;12:369–76.
Dame JB, Murrel D, Worley DE, Schad GA. Trichinella spiralis: genetic evidence for synanthropic subspecies in sylvatic hosts. Exp Parasitol. 1987;64:195–203.
La Rosa G, Pozio E, Rossi P. Biochemical resolution of European and African isolates of Trichinella nelsoni Britov and Boev, 1972. Parasitol Res. 1990;77:173–6.
La Rosa G, Pozio E, Rossi P, Murrel KD. Allozyme analysis of Trichinella isolates from various host species and geographical regions. J Parasitol. 1992;78:641–6.
La Rosa G, Pozio E. Molecular investigation of African isolates of Trichinella reveals genetic polymorphism in Trichinella nelsoni. Int J Parasitol. 2000;(5):663–7.
La Rosa G, Marucci G, Zarlenga DS, Pozio E. Trichinella pseudospiralis populations of the Palearctic region and their relationship with populations of the Nearctic and Australian regions. Int J Parasitol. 2001;31:297–305.
La Rosa G, Marucci G, Zarlenga DS, Casulli A, Zarnke RL, Pozio R. Molecular identification of natural hybrids between Trichinella nativa and Trichinella T6 provides evidence of gene flow and ongoing genetic divergence. Int J Parasitol. 2003;33:209–16.
Zarlenga DS, Aschenbrebber RA, Lechtenfels JR. Variations in microsatellite sequences provide evidence for population differences and multiple ribosomal gene repeats in Trichinella pseudospiralis. J Parasitol. 1996;82:534–8.
Otranto D, Testini G, De Luca F, Hu M, Shamsi S, Gasser RB. Analysis of genetic variability within Thelazia callipaeda (Nematoda: Thelazioidea) from Europe and Asia by sequencing and mutation scanning of the mitochondrial cytochrome c oxidase subunit 1 gene. Mol Cel Probe. 2005;19:306–13.
Di Cisare A, Otranto D, Latrofa MS, Veronisi F, Perrucci S, Lalosevic D, et al. Genetic variability of Eucoleus aerophilus from domestic and wild hosts. Res Vet Sci. 2014;96:512–5.
Rothmann W, de Waal PJ. Diversity of Spirocerca lupi in domestic dogs and black-backed jackals (Canis mesomelas) from South Africa. Vet Parasitol. 2017;244:59–63.
Berntzen AK. Comparative growth and development of Trichinella spiralis in vitro and in vivo, with a redescription of the life cycle. Exp Parasitol. 1965;16:74–106.
Montgomery WI, Wilson WL, Hamilton R, McCartney P. Dispersion in the wood mouse, Apodemus sylvaticus: variable resources in time and space. J Anim Ecol. 1991;60:179–92.
Mikesic DG, Drickamer LC. Factors affecting home-range size in house mice (Mus musculus domesticus) living in outdoor enclosures. Am Midl Nat. 1992;127:31–40.
Pocock JO, Hauffe HC, Searle JB. Dispersal in house mice. Biol J Linn Soc. 2005;84:565–83.
Gardner-Santana LC, Norris DE, Fornadel CE, Hinson ER, Klein SL, Glass GE. Commensal ecology, urban landscapes, and their influence on the genetic characteristics of city-dwelling Norway rats (Rattus norvegicus). Mol Ecol. 2009;18:2766–78.
Nieberding C, Libois R, Douady CJ, Morand S, Michaux JR. Phylogeography of a nematode (Heligmosomoides polygyrus) in the western Palearctic region: persistence of northern cryptic populations during ice ages? Mol Ecol. 2005;14:765–79.
Nieberding C, Morand S, Libois R, Michaux JR. Parasites and the island syndrome: the colonization of the western Mediterranean islands by Heligmosomoides polygyrus (Dujardin, 1845). J Biogeogr. 2006;33:1212–22.
Nieberding C, Durette-Desset M-C, Vanderpoorten A, Casanova JC, Ribas A, Deffontaine V, et al. Geography and host biogeography matter for understanding the phylogeography of a parasite. Mol Phylogenet Evol. 2008;47:538–54.
Monroy FG, Enriquez FJ. Heligmosomoides polygyrus: a model for chronic gastrointestinal helminthiasis. Parasitol Today. 1992;8:49–54.
Callejón R, de Rojas M, Nieberding C, Foronda P, Feliú C, Guevera D, Cutillas C. Molecular evolution of Trichuris muris isolated from different Muridae hosts in Europe. Parasitol Res. 2010;107:631–41.
Callejón R, de Rojas M, Feliú C, Balao F, Marrugal A, Henttonen H, et al. Phylogeography of Trichuris populations isolated from different Cricetidae rodents. Parasitology. 2012;139:1795–812.
Wasimuddin BJ, Ribas A, Baird SJE, Piálek J, Goüy de Bellocq J. Testing parasite ‘intimacy’: the whipworm Trichuris muris in the European house mouse hybrid zone. Ecol Evol. 2016;6:2688–701.
Fahmy MAM. An investigation on the life cycle of Trichuris muris. Parasitology. 1954;44:50–7.
Gregory RD, Keymer AE, Clarge JR. Genetics, sex and exposure: the ecology of Heligmosomoides polygyrus (Nematoda) in the wood mouse. J Anim Ecol. 1990;59:363–78.
Yong HS, Eamsobhana P. Definitive rodent hosts of the rat lungworm Angiostrongylus cantonensis. Raffles B Zool. 2013;29:111–5.
Liu CY, Zhang RL, Chen MX, Li J, Ai L, Wu CY, et al. Characterisation of Angiostrongylus cantonensis isolates from China by sequences of internal transcribed spacers of nuclear ribosomal DNA. J Anim Vet Adv. 2011;10:593–6.
Thaenkham U, Pakdee W, Nuamtanong S, Maipannich W, Pubampen S, Sa-Nguankiat S, Komalamisra C. Population structure of Angiostrongylus cantonensis (Nematoda: Metastrongylidae) in Thailand based on PCR-RAPD markers. SE Asian J Trop Med. 2012;43:567–73.
Dusitsittipon S, Thaenkham D, Watthanakulpanich D, Adisakwattana P, Komalamisra C. Genetic differences in the rat lungworm, Angiostrongylus cantonensis (Nematoda: Angiostrongylidae), in Thailand. J Helminthol. 2015;89:545–51.
Yong H-S, Eamsobhana P, Song S-L, Prasartvit A, Lim P-E. Molecular phylogeography of Angiostrongylus cantonensis (Nematoda: Angiostrongylidae) and genetic relationships with congeners using cytochrome b gene marker. Acta Trop. 2015;148:66–71.
Vitta A, Srisongcram N, Thiproaj J, Wongma A, Polsut W, Fukrusa C, et al. Phylogeny of Angiostrongylus cantonensis in Thailand based on cytochrome c oxidase subunit I gene sequence. SE Asian J Trop Med. 2016;47:377–86.
Eamsobhana P, Song S-L, Yong H-S, Prasartvit A, Boonyong S, Tungtrongchitr A. Cytochrome c oxidase subunit I haplotype diversity of Angiostrongylus cantonensis (Nematoda: Angiostrongylidae). Acta Trop. 2017;171:141–5.
Tokiwa T, Harunari T, Tanikawa T, Komatsu N, Koizumi N, Tung K-C, et al. Phylogenetic relationships of rat lungworm, Angiostrongylus cantonensis, isolated from different geographical regions revealed widespread multiple lineages. Parasitol Int. 2012;61:431–6.
Monte TCC, Simões RO, Oliveira APM, Novaes CF, Thiengo SC, Silva AJ, et al. Phylogenetic relationship of the Brazilian isolates of the rat lungworm Angiostrongylus cantonensis (Nematoda: Metastrongylidae) employing mitochondrial COI gene sequence data. Parasit Vectors. 2012;5:248.
Eamsobhana P, Yong HS, Song SL, Prasartvit A, Boonyong S, Tungtongchitr A. Cytochrome c oxidase subunit I haplotype reveals high genetic diversity of Angiostrongylus malaysiensis (Nematoda: Angiostrongylidae). J Helminthol. 2018;92:254–9.
Rodpai R, Intapan PM, Thanchomnang T, Sanpool O, Sadaow L, Laymanivong S, et al. Angiostrongylus cantonensis and A. malaysiensis broadly overlap in Thailand, Lao PDR, Cambodia and Myanmar: a molecular survey of larvae in land snails. PLoS One. 2016;11:e0161128.
Yong HS, Song SL, Eamsobhana P, Goh SY, Lim PE. Complete mitochondrial genome reveals genetic diversity of Angiostrongylus cantonensis (Nematoda: Angiostrongylidae). Acta Trop. 2015;152:157–64.
Dusitsittipon S, Criscione CD, Morand S, Komalamisra C, Thaenkham U. Cryptic lineage diversity in the zoonotic pathogen Angiostrongylus cantonensis. Mol Phylogenet Evol. 2017;107:404–14.
Dusitsittipon S, Criscione CD, Morand S, Komalamisra C, Thaenkham U. Hurdles in the evolutionary epidemiology of Angiostrongylus cantonensis: pseudogenes, incongruence between taxonomy and DNA sequence variants, and cryptic lineages. Evol Appl. 2018;11:1257–69.
Brouat C, Tatard C, Machin A, Kane M, Diouf M, Bâ K, Duplantier JM. Comparative population genetics of a parasitic nematode and its host community: the trichostrongylid Neoheligmonella granjoni and Mastomys rodents in south-eastern Senegal. Int J Parasitol. 2011;41:1301–9.
Brouat C, Loiseau A, Kane M, Bâ K, Duplantier J-M. Population genetic structure of two ecologically distinct multimammate rats: the commensal Mastomys natalensis and the wild M. erythroleucus in south-eastern Senegal. Mol Ecol. 2007;16:2985–97.
Brant SV, Ortí G. Evidence for gene flow in parasitic nematodes between two host species of shrews. Mol Ecol. 2003;12:2853–9.
Fisher MC, Viney ME. The population genetic structure of the facultatively sexual parasitic nematode Strongyloides ratti in wild rats. Proc R Soc B Biol Sci. 1998;265:703–9.
Viney ME. A genetic analysis of reproduction in Strongyloides ratti. Parasitology. 1994;109:511–5.
Paterson S, Fisher MC, Viney ME. Inferring infection processes of a parasitic nematode using population genetics. Parasitology. 2000;120:185–94.
Müller-Graf CD, Durand P, Feliu C, Hugot J-P, O’Callaghan CJ, Renaud F, et al. Epidemiology and genetic variability of two species of nematodes (Heligmosomoides polygyrus and Syphacia stroma) of Apodemus spp. Parasitology. 1999;118:425–32.
Morand S. Life-history traits in parasitic nematodes: a comparative approach for the search of invariants. Funct Ecol. 1996;10:210–8.
Dallas JF, Irvine RJ, Halvorsen O. DNA evidence that Ostertagia gruehneri and Ostertagia arctica (Nematoda: Ostertagiinae) in reindeer from Norway and Svalbard are conspecific. Int J Parasitol. 2000;30:655–8.
Dallas JF, Irvine RJ, Halvorsen O. DNA evidence that Marshallagia marshalli Ransom, 1907 and M. occidentalis Ransom, 1907 (Nematoda: Ostertagiinae) from Svalbard reindeer are conspecific. Syst Parasitol. 2001;50:101–3.
Hoberg EP, Monsen KJ, Kutz S, Blouin MS. Structure, biodiversity, and historical biogeography of nematode faunas in Holarctic ruminants: morphological and molecular diagnoses for Teladorsagia boreoarticus n. sp. (Nemadota: Ostertagiinae), dimorphic cryptic species in muskoxen (Ovibos moschatus). J Parasitol. 1999;85:910–34.
Blouin MS, Yowell CA, Courtney CH, Dame JB. Host movement and the genetic structure of populations of parasitic nematodes. Genetics. 1995;141:1007–14.
Tyler NJC, Øritsland NA. Home ranges in Svalbard reindeer. Rangifer. 1990;S3:147–8.
Long ES, Diefenbach DR, Rosenberry CS, Wallingford BD. Multiple proximate and ultimate causes of natal dispersal in white-tailed deer. Behav Ecol. 2008;19:1235–42.
Ács Z, Hayward A, Sugár L. Genetic diversity and population genetics of large lungworms (Dictyocaulus, Nematoda) in wild deer in Hungary. Parasitol Res. 2016;115:3295–312.
Archie EA, Ezenwa VO. Population genetic structure and history of a generalist parasite infecting multiple sympatric host species. Int J Parasitol. 2011;41:89–98.
Falk BG, Perkins SL. Host specificity shapes population structure of pinworm parasites in Caribbean reptiles. Mol Ecol. 2013;22:4576–90.
Calsbeek R. Sex-specific adult dispersal and its selective consequences in the brown anole, Anolis sagrei. J Anim Ecol. 2009;78:617–24.
Jorge F, Roca V, Perera A, Harris DJ, Carretero MA. A phylogenetic assessment of the colonisation patterns in Spauligodon atlanticus Astasio-Arbiza et al., 1987 (Nematoda: Oxyurida: Pharyngodonidae), a parasite of lizards of the genus Gallotia Boulenger: no simple answers. Syst Parasitol. 2011;80:53–66.
Gustafson KD, Newman RA, Rhen T, Tkach VV. Spatial and genetic structure of directly-transmitted parasites reflects the distribution of their specific amphibian hosts. Pop Ecol. 2018;60:261–73.
Johnson PCD, Webster LMI, Adam A, Buckland R, Dawson DA, Keller LF. Abundant variation in microsatellites of the parasitic nematode Trichostrongylus tenuis and linkage to a tandem repeat. Mol Biochem Parasit. 2006;148:210–8.
Webster LMI, Johnson PCD, Adam A, Mable BK, Keller MF. Macrogeographic population structure in a parasitic nematode with avian hosts. Vet Parasitol. 2007;144:93–103.
Seivwright LJ, Redpath SM, Mougeot F, Watt L, Hudson PJ. Faecal egg counts provide a reliable measure of Trichostrongylus tenuis intensities in free-living red grouse Lagopus lagopus scoticus. J Helminthol. 2004;78:69–76.
Wernham CV, Toms MP, Marchant JH, Clark JA, Siriwardena GM, Baillie SE. Migration Atlas: Movements of the Birds of Britain and Ireland. London: Poyser; 2002.
Nagasawa K. The life cycle of Anisakis simplex: a review. In: Ishikura H, Kikuchi K, editors. Intestinal Anisakiasis in Japan. Tokyo: Springer; 1990. p. 31–40.
Åkesson S, Weimerskirch H. Albatross long-distance navigation: comparing adults and juveniles. J Navigation. 2005;58:365–73.
Tsai Y-JJ, Mann J. Dispersal, philopatry, and the role of fission-fusion dynamics in bottlenose dolphins. Mar Mammal Sci. 2012;29:261–79.
Mattiucci S, Nascetti G, Cianchi R, Paggi L, Arduino P, Margolis L, et al. Genetic and ecological data on the Anisakis simplex complex, with evidence for a new species (Nematoda, Ascaridoidea, Anisakidae). J Parasitol. 1997;83:401–16.
Mattiucci S, Nascetti G, Dailey M, Webb SC, Barros NB, Cianchi R, Bullini L. Evidence for a new species of Anisakis Dujardin, 1845: morphological description and genetic relationships between congeners (Nematoda: Anisakidae). Syst Parasitol. 2005;61:157–71.
Pontes T, D’Amelio S, Costa G, Paggi L. Molecular characterization of larval anisakid nematodes from marine fishes of Madeira by a PCR-based approach, with evidence for a new species. J Parasitol. 2005;91:1430–4.
Beverly-Burton M. Population genetics of Anisakis simplex (Nematoda: Ascaridoidea) in Atlantic salmon (Salmo salar) and their use as biological indicators of host stocks. Environ Biol Fish. 1978;3:369–77.
Mattiucci S, Nascetti G. Genetic diversity and infection levels of anisakid nematodes parasitic in fish and marine mammals from Boreal and Austral hemispheres. Vet Parasitol. 2007;148:43–57.
Lyons ET, Spraker TR, de Long RL, Ionita M, Melin SR, Nadler SA, Tolliver SC. Review of research on hookworms (Uncinaria lucasi Stiles, 1901) in northern fur seals (Callorhinus ursinus Linnaeus, 1758). Parasitol Res. 2011;109:257–65.
Campbell RA, Gales NJ, Lento GM, Baker CS. Islands in the sea: extreme female natal site fidelity in the Australian sea lion, Neophoca cinerea. Biol Lett. 2008;4:139–42.
Haynes BT, Marcus AD, Higgins DP, Gongora J, Gray R, Šlapeta J. Unexpected absence of genetic separation of a highly diverse population of hookworms from geographically isolated hosts. Infect Genet Evol. 2014;28:192–200.
Nadler SA, Adams BJ, Lyons ET, De Long RL, Melin SR. Molecular and morphometric evidence for separate species of Uncinaria (Nematoda: Ancylostomatidae) in California sea lions and northern fur seals: hypothesis testing supplants verification. J Parasitol. 2000;86:1099–106.
Martín-Sánchez J, Paniagua I, Valero A. Contribution to the knowledge of Hysterothylacium aduncum through electrophoresis of the enzymes glucose phosphate isomerase and phosphoglucomutase. Parasitol Res. 1998;84:160–3.
Klimpel S, Kleinertz S, Hanel R, Rükert S. Genetic variability in Hysterothylacium aduncum, a raphidascarid nematode isolated from sprat (Sprattus sprattus) of different geographical areas of the northeastern Atlantic. Parasitol Res. 2007;101:1425–30.
Limborg MT, Pedersen JS, Hemmer-Hansen J, Tomkeiwicz J, Bekkevold D. Genetic population structure of European sprat Sprattus sprattus: differentiation across a steep environmental gradient in a small pelagic fish. Mar Ecol. 2009;379:213–24.
Martín-Sánchez J, Díaz M, Artacho ME, Valero A. Molecular arguments for considering Hysterothylacium fabri (Nematoda: Anisakidae) a complex of sibling species. Parasitol Res. 2003;89:214–20.
Mejía-Madrid HH, Vázquez-Domínguez E, Pérez-Ponce de León G. Phylogeography and freshwater basins in central Mexico: recent history as revealed by the fish parasite Rhabdochona lichtenfelsi (Nematoda). 2007;34:787–801.
Li WX, Wang GT, Nie P. Genetic variation of fish parasite populations in historically connected habitats: undetected habitat fragmentation effect on populations of the nematode Procamallanus fulvidraconis in the catfish Pelteobagrus fulvidraco. J Parasitol. 2008;94:643–7.
Ma R, Yang G, Duan H, Jiang J, Wang S, Feng X. China’s lakes at present: number, area and spatial distribution. Sci China Earth Sci. 2011;54:283–9.
Wu SG, Wang GT, Xi BW, Xiong F, Liu T, Nie P. Population genetic structure of the parasitic nematode Camallanus cotti inferred from DNA sequences of ITS1 rDNA and the mitochondrial COI gene. Vet Parasitol. 2009;164:248–56.
Blouin MS, Liu J, Berry RE. Life cycle variation and the genetic structure of nematode populations. Heredity. 1999;83:253–9.
Johnigk S-A, Ehlers R-U. Juvenile development and life cycle of Heterorhabditis bacteriophora and H. indica (Nematoda: Heterorhabditidae). Nematology. 1999;1:251–60.
Belaich MN, Buldain D, Ghiringhelli PD, Hyman B, Micieli MV, Achinelly MF. Nucleotide sequence differentiation of Argentine isolates of the mosquito parasitic nematode Strelkovimermis spiculatus (Nematoda: Mermithidae). 2015;40:415–8.
Crainey JL, Wilson MD, Post RJ. An 18S ribosomal DNA barcode for the study of Isomermis lairdi, a parasite of the blackfly Simulium damnosum s.l. Met Vet Entomol. 2009;23:238–44.
Tobias ZJ, Jorge F, Poulin R. Life at the beach: comparative phylogeography of a sandhopper and its nematode parasite reveals extreme lack of parasite mtDNA variation. Biol J Linn Soc. 2017;122:113–32.
Tang S, Hyman BC. Mitochondrial genome haplotype hypervariation within the isopod parasitic nematode Thaumamermis cosgrovei. Genetics. 2007;176:1139–50.
Jobet E, Durand P, Langand J, Müller-Graf CDM, Hugot J-P, Bounnoux M-E, et al. Comparative genetic diversity of parasites and their hosts: population structure of an urban cockroach and its haplo-diploid parasite (oxyuroid nematode). Mol Ecol. 2000;9:481–6.
Morand S, Rivault C. Infestation dynamics of Blatticola blattae Graeffe (Nematoda: Thelastomatidae), a parasite of Blattella germanica (Dictyoptera: Blattellidae). Int J Parasitol. 1992;22:983–9.
Lin Q, Li HM, Gao M, Wang XY, Ren WX, Cong MM, et al. Characterization of Baylisascaris schroederi from Qinling subspecies of giant panda in China by the first internal transcribed spacer (ITS-1) of nuclear ribosomal DNA. Parasitol Res. 2012;110:1297–303.
Zhao GH, Li HM, Ryan UM, Cong MM, Hu B, Gao M, et al. Phylogenetic study of Baylisascaris schroederi isolated from Qinling subspecies of giant panda in China based on combined nuclear 5.8S and the second internal transcribed spacer (ITS-2) ribosomal DNA sequences. Parasitol Int. 2012;61:497–500.
Zhou X, Xie Y, Zhang Z-H, Wang C-D, Sun Y, Gu X-B, et al. Analysis of the genetic diversity of the nematode parasite Baylisascaris schroederi from wild giant pandas in different mountain ranges in China. Parasit Vectors. 2013;6:233.
Xie Y, Zhou X, Zhang Z, Wang C, Sun Y, Liu T, et al. Absence of genetic structure in Baylisascaris schroederi populations, a giant panda parasite, determined by mitochondrial sequencing. Parasit Vectors. 2014;7:606.
Lu Z, Johnson WE, Menotti-Raymong M, Yuhki N, Martenson JS, Mainka S, et al. Patterns of genetic diversity in remaining giant panda populations. Conserv Biol. 2002;15:1596–607.
Peng Z, Zhang C, Shen M, Bao H, Hou Z, He S, et al. Baylisascaris schroederi infection in giant pandas (Ailuropoda melanoleuca) in Foping National Nature Reserve, China. J Wildlife Dis. 2017;53:854–8.
Wei F, Hu Y, Zhu L, Bruford MW, Zhan X, Zhang L. Black and white and read all over: the past, present and future of giant panda genetics. Mol Ecol. 2012;21:5660–74.
Solórzano-García B, Ellis EA, Rodríguez-Luna E. Deforestation and primate habitat availability in Los Tuxtlas Biosphere Reserve Mexico. Int J Ecosystem. 2012;2:61–6.
Solórzano-García B, Gasca-Pineda J, Poulin R, Pérez-Ponce de León G. Lack of genetic structure in pinworm populations from New World primates in forest fragments. Int J Parasitol. 2017;47:941–50.
Gasser RB, Woods WG, Blotkamp C, Verweij JJ, Storey PA, Polderman AM. Screening for nucleotide variations in ribosomal DNA arrays of Oesophagostomum bifurcum by polymerase chain reaction-coupled single-strand conformation polymorphism. Electrophoresis. 1999;20:1486–91.
De Gruijter JM, Ziem J, Verweij JJ, Polderman AM, Gasser RB. Genetic substructuring within Oesophagostomum bifurcum (Nematoda) from human and non-human primates from Ghana based on random amplified polymorphic DNA analysis. Am J Trop Med Hyg. 2004;71:227–33.
Gasser RB, de Gruijter JM, Polderman AM. Insights into the epidemiology and genetic make-up of Oesophagostomum bifurcum from human and non-human primates using molecular tools. Parasitology. 2006;132:453–60.
De Gruijter JM, Gasser RB, Polderman AM, Asigri V, Dijkshoorn L. High resolution DNA fingerprinting by AFLP to study the genetic variation among Oesophagostomum bifurcum (Nematoda) from human and non-human primates from Ghana. Parasitology. 2005;130:229–37.
Labes EM, Wijayanti N, Deplazes P, Mathis A. Genetic characterization of Strongyloides spp. from captive, semi-captive and wild Bornean orangutans (Pongo pygmaeus) in Central and East Kalimantan, Borneo, Indonesia. Parasitology. 2011;138:1417–22.
Ravasi DF, O’Riain MJ, Davids F, Illing N. Phylogenetic evidence that two distinct Trichuris genotypes infect both humans and non-human primates. PLoS One. 2012;7:e44187.
Makouloutou P, Mbehang Nguema PP, Fujita S, Takenoshita Y, Hasegawa H, Yanagida T, Sato H. Prevalence and genetic diversity of Oesophagostomum stephanostomum in wild lowland gorillas at Moukalaba-Doudou National Park, Gabon. Helminthologia. 2014;51:83–93.
Blizzard EL, David CD, Henke S, Long DB, Hall CA, Yabsley MJ. Distribution, prevalence, and genetic characterization of Baylisascaris procyonis in selected areas of Georgia. J Parasitol. 2010;96:1128–33.
Sarkissian CA, Campbell SK, Dharmarajan G, Jacquot J, Page LK, Graham DH. Microgeographic population genetic structure of Baylisascaris procyonis (Nematoda: Ascaroidae) in western Michigan indicates the Grand River is a barrier to gene flow. J Parasitol. 2015;101:671–6.
Osten-Sacken N, Heddegrott M, Schleimer A, Anheyer-Behmenburg HE, Runge M, Horsburgh GJ, et al. Similar yet different: co-analysis of the genetic diversity and structure of an invasive nematode parasite and its invasive mammalian host. Int J Parasitol. 2018;48:233–43.
Dubey S, Shine R. Origin of the parasites of an invading species, the Australian cane toad (Bufo marinus): are the lungworms Australian or American? Mol Ecol. 2008;17:4418–24.
Sheng L, Cui P, Fang S-F, Lin R-Q, Zou F-C, Zhu X-Q. Sequence variability in four mitochondrial genes among rabbit pinworm (Passalurus ambiguus) isolates from different localities in China. Mitochondr DNA. 2015;26:501–4.
Mayr E. Animal Species and Evolution. Cambridge: Harvard University Press; 1963.
Gaither MR, Abey G, Vignon M, Meguro Y-I, Rigby M, Runyon C, et al. An invasive fish and the time-lagged spread of Its parasite across the Hawaiian Archipelago. PLoS One. 2013;8:e56940.
Gagne RB, Sprehn CG, Alda F, McIntyre PB, Gilliam JF, Blum MJ. Invasion of the Hawaiian Islands by a parasite infecting imperiled stream fishes. Ecography. 2018;41:528–39.
Knopf K, Mahnke M. Differences in susceptibility of the European eel (Anguilla anguilla) and the Japanese eel (Anguilla japonica) to the swim-bladder nematode Anguillicola crassus. Parasitology. 2004;129:491–6.
Rahhou I, Morand S, Lecomte-Finiger R, Sasal P. Biogeographical relationships of the eel parasite Anguillicola crassus revealed by random amplified polymorphic DNA markers (RAPDS). B Fr Pêch Piscic. 2005;378-379:87–98.
Wielgoss S, Tarachewski H, Meyer A, Wirth T. Population structure of the parasitic nematode Anguillicola crassus, an invader of declining North Atlantic eel stocks. Mol Ecol. 2008;17:3478–95.
Wielgoss S, Holland F, Wirth T, Meyer A. Genetic signatures in an invasive parasite of Anguilla anguilla correlate with differential stock management. J Fish Biol. 2010;77:191–210.
Wit J, Gilleard JS. Resequencing helminth genomes for population and genetic studies. Trends Parasitol. 2017;33:388–99.
Korhonen PK, Young ND, Gasser RB. Making sense of genomes of parasitic worms: tackling bioinformatic challenges. Biotechnol Adv. 2016;34:663–86.
Baird NA, Etter PD, Atwood TS, Currey MS, Shiver AL, Lewis ZA, et al. Rapid SNP discovery and genetic mapping using sequenced RAD markers. PLoS One. 2008;3e:3376.
Peterson BK, Webster JN, Kay EH, Fisher HS, Hoekstra HE. Double digest RADseq: an inexpensive method for de novo SNP discovery and genotyping in model and non-model species. PLoS One. 2012;7:e37135.
Da Fonseca RR, Albrechtsen A, Espregueira Themudo G, Ramos-Madrigal J, Andreas Sibbesen J, Maretty L, et al. Next-generation biology: sequencing and data analysis approaches for non-model organisms. Mar Genom. 2016;30:3–13.
Puckett EE, Park J, Combs M, Blum MJ, Bryant JE, Caccone A, et al. Global population divergence and admixture of the brown rat (Rattus norvegicus). Proc R Soc B Biol Sci. 2016;283:20161762.
Anderson EC, Gerke JP, Shapiro JA, Crissman JR, Ghosh R, Bloom JS, et al. Chromosome-scale selective sweeps shape Caenorhabditis elegans genomic diversity. Nat Genet. 2012;44:285–90.
Li H, Durbin R. Inference of human population history from individual whole-genome sequences. Nature. 2011;475:493–6.
Jones FC, Grabherr MG, Chan YF, Russel P, Mauceli E, Johnson J, et al. The genomic basis of adaptive evolution in threespine sticklebacks. Nature. 2012;484:55–61.
Ott J, Wang J, Leal SM. Genetic linkage analysis in the age of whole-genome sequencing. Nat Rev Genet. 2015;16:275–84.
Fumagalli M. Assessing the effect of sequencing depth and sample size in population genetics inferences. PLoS One. 2013;8:e79667.
Hosono S, Faruqi AF, Dean FB, Du Y, Sun Z, Wu X, et al. Unbiased whole-genome amplification directly from clinical samples. Genome Res. 2003;13:954–64.
Sabina J, Leamon JH. Bias in whole genome amplification: causes and considerations. Methods Mol Biol. 2015;1347:15–41.
Mobegi VA, Duffy CW, Amambua-Ngwa A, Loua KM, Laman E, Nwakanma DS, et al. Genome-wide analysis of selection on the malaria parasite Plasmodium falciparum in West African populations of differing infection endemicity. Mol Biol Evol. 2014;31:4190–9.
Coles GC, Jackson F, Pomroy WE, Prichard RK, von Samson-Himmelstjerna G, Silvestre A, et al. The detection of anthelmintic resistance in nematodes of veterinary importance. Vet Parasitol. 2006;136:167–85.
Choi Y-J, Bisset SA, Doyle SR, Hallsworth-Pepin K, Martin J, Grant WN, Mitreva M. Genomic introgression mapping of field-derived multiple-anthelmintic resistance in Teladorsagia circumcincta. PLoS Genet. 2017;13:e1006857.
Small ST, Reimer LJ, Tisch DJ, King CL, Christensen BM, Siba PM, et al. Population genomics of the filarial nematode parasite Wuchereria bancrofti from mosquitoes. Mol Ecol. 2016;25:1465–77.
Kikuchi T, Hino A, Tanaka T, Aung MPPTHH, Afrin T, Nagayasu E, et al. Genome-wide analyses of individual Strongyloides stercoralis (Nematoda: Rhabditoidea) provide insights into population structure and reproductive life cycles. PLoS Negl Trop Dis. 2016;10:e0005253.
Gendron St-Marseille A-F, Lord E, Véronneau P-Y, Brodeur J, Mimee B. Genome scans reveal homogenization and local adaptations in populations of the soybean cyst nematode. Front Plant Sci. 2018;9:987.
Lu F, Lipka AE, Glaubitz J, Elshire R, Cherney JH, Casler MD, et al. Switchgrass genomic diversity, ploidy, and evolution: novel insights from a network-based SNP discovery protocol. PLoS Genet. 2013;9:e1003215.
Chartier C, Reche B. Gastrointestinal helminths and lungworms of French dairy goats: prevalence and geographical distribution in Poitou-Charentes. Vet Res Commun. 1992;16:327–35.
Kataman KK, Thamsborg SM, Dalsgaard A, Kyvsgaard NC, Mejer H. Environmental contamination and transmission of Ascaris suum in Danish organic pig farms. Parasit Vectors. 2016;9:80.
Shokralla S, Spall JL, Gibson JF, Hadibabaei M. Next-generation sequencing technologies for environmental DNA research. Mol Ecol. 2012;21:1794–805.
Peham T, Steiner FM, Schlick-Steiner BC, Arthofer W. Are we ready to detect nematode diversity by next generation sequencing? Ecol Evol. 2017;7:4147–51.
Wimmer B, Craig BH, Pilkington JG, Pemberton JM. Non-invasive assessment of parasitic nematode species diversity in wild Soay sheep using molecular markers. Int J Parasitol. 2004;34:625–31.
Blouin MS, Dame JB, Tarrant CA, Courtney CH. Unusual population genetics of a parasitic nematode: mtDNA variation within and among populations. Evolution. 1992;46:470–6.
Höglund J, Engström A, Morrison DA, Mattsson JC. Genetic diversity assessed by amplified fragment length polymorphism analysis of the parasitic nematode Dictyocaulus viviparus the lungworm of cattle. Int J Parasitol. 2004;34:475–84.
Banks DJD, Singh R, Barger IA, Pratap B, le Jambre LF. Development and survival of infective larvae of Haemonchus contortus and Trichostrongylus colubriformis on pasture in a tropical environment. Int J Parasitol. 1990;20:155–60.
Wright S. Evolution in Mendelian populations. Genetics. 1931;16:97–159.
Slatkin M. Gene flow in natural populations. Ann Rev Ecol Syst. 1985;16:393–430.
Stern C. The Hardy-Weinberg law. Science. 1943;97:137–8.
Andrews RH, Chilton NB. Multilocus enzyme electrophoresis: a valuable technique for providing answers to problems in parasite systematics. Int J Parasitol. 1999;29:213–53.
Slatkin M. Linkage disequilibrium - understanding the evolutionary past and mapping the medical future. Nat Genet. 2008;9:477–85.
Avise JC. Phylogeography: The History and Formation of Species. Cambridge: Harvard University Press; 2000.
Hartl DL. A primer of population genetics. Massachussets: Sinauer; 1988.
Lynch M, Mulligan BG. Analysis of population genetic structure with RAPD markers. Mol Ecol. 1994;3:91–9.
Powers TO, Todd TC, Burnell AM, Murray PCB, Fleming CC, Szalanski AL, et al. The rDNA internal transcribed spacer region as a taxonomic marker for nematodes. J Nematol. 1997;29:441–50.
McVean G. The structure of linkage disequilibrium around a selective sweep. Genetics. 2007:1395–406.
Obendorf DL, Beveridge I, Andrews RH. Cryptic species in populations of Globocephaloides trifidospicularis Kung (Nematoda: Trichostrongyloidea), parasitic in macropodid marsupials. Trans Roy Soc South Aus. 1991;115:213–6.
Jabbar A, Beveridge I, Mohandas N, Chilton NB, Littlewood DTJ, Jex AR, Gasser RB. Analyses of mitochondrial amino acid sequence datasets support the proposal that specimens of Hypodontus macropi from three species of macropodid hosts represent distinct species. BMC Evol Biol. 2013;13:259.
Chilton NB, Beveridge I, Andrews RH. Electrophoretic comparison of Rugopharynx longibursaris Kung and R. omega Beveridge (Nematoda: Strongyloidea), with the description of R. sigma n. sp. from pademelons, Thylogale spp. (Marsupialia: Macropodidae). Syst Parasitol. 1992;26:159–169.
Chilton NB, Beveridge I, Hoste H, Gasser RB. Evidence for hybridisation between Paramacropostrongylus iugalis and P. typicus (Nematoda: Strongyloidea) in grey kangaroos, Macropus fuliginosus and M. giganteus, in a zone of sympatry in eastern Australia. Int J Parasitol. 1997;27:475–82.
Chilton NB, Beveridge I, Andrews RH. Electrophoretic and morphological analysis of Paramacropostrongylus typicus (Nematoda: Strongyloidea), with the description of a new species, Paramacropostrongylus iugalis, from the eastern grey kangaroo Macropus giganteus. Syst Parasitol. 1993;24:35–44.
Beveridge I, Chilton NB, Andrews RH. Sibling species within Macropostrongyloides baylisi (Nematoda: Strongyloidea) from macropodid marsupials. Int J Parasitol. 1993;23:21–33.
Beveridge I, Chilton NB, Andrews RH. A morphological and electrophoretic study of Rugopharynx zeta (Johnston & Mawson, 1939) (Nematoda: Strongyloidea), with the description of a new species, R. mawsonae, from the black-striped wallaby Macropus dorsalis (Marsupialia: Macropodidae). Syst Parasitol. 1994;27:159–71.
Chilton NB, Smales LR. An electrophoretic and morphological analysis of Labiostrongylus (Labiomultiplex) uncinatus (Nematoda: Cloacinidae), with the description of a new species, L. contiguus, from Macropus parryi (Marsupialia: Macropodidae). Syst Parasitol. 1996;35:49–57.
Smales LR, Chilton NB. An electrophoretic and morphological analysis of Labiostrongylus (Labiosimplex) bancrofti (Johnston & Mawson, 1939) (Nematoda: Cloacinidae), from macropodid marsupials, with the description of two new species. Syst Parasitol. 1997;36:193–201.
Huby-Chilton F, Beveridge I, Gasser RB, Chilton NB. Redescription of Zoniolaimus mawsonae Beveridge, 1983 (Nematoda: Strongyloidea) and the description of Z. latebrosus n. sp. from the red kangaroo Macropus rufus (Marsupialia: Macropodidae) based on morphological and molecular data. Syst Parasitol. 2002;51:135–47.
Chilton NB, Huby-Chilton F, Gasser RB, Beveridge I. Review of Papillostrongylus Johnston & Mawson, 1939 (Nematoda: Strongyloidea) from wallabies and kangaroos (Marsupialia: Macropodidae) using morphological and molecular techniques, with the description of P. barbatus n. sp. Syst Parasitol. 2002;51:81–93.
Nascetti G, Paggi L, Orecchia P, Smith JW, Mattiucci S, Bullini L. Electrophoretic studies on the Anisakis simplex complex (Ascaridida: Anisakidae) from the Mediterranean and North-East Atlantic. Int J Parasitol. 1986;16:633–40.
Mattiucci S, Paggi L, Nascetti D, Portes Santos C, Costa G, di Beneditto AP, et al. Genetic markers in the study of Anisakis typica (Diesing, 1860): larval identification and genetic relationships with other species of Anisakis Dujardin, 1845 (Nematoda: Anisakidae). Syst Parasitol. 2002;51:159–70.
Abollo E, Paggi L, Pascual S, D’Amelio S. Occurrence of recombinant genotypes of Anisakis simplex s.s. and Anisakis pegreffii (Nematoda: Anisakidae) in an area of sympatry. Infect Genet Evol. 2003;3:175–81.
Martín-Sánchez J, Artacho-Reinoso ME, Díaz-Gavilán M, Valero-López A. Structure of Anisakis simplex s.l. populations in a region sympatric for A. pegreffii and A. simplex s.s.: absence of reproductive isolation between both species. Mol Biochem Parasitol. 2005;141:155–62.
Klimpel S, Busch MW, Kuhn T, Rhode A, Palm FW. The Anisakis simplex complex off the South Shetland Islands (Antarctica): endemic populations versus introduction through migratory hosts. Mar Ecol Prog Ser. 2010;403:1–11.
Baldwin RE, Rew MB, Johansson ML, Banks MA, Jacobson KC. Population structure of three species of Anisakis nematodes recovered from Pacific sardines (Sardinops sagax) distributed throughout the California current system. J Parasitol. 2011;97:545–54.
Cross MA, Collins C, Campbell N, Watts PC, Chubb JC, Cunningham CO, et al. Levels of intra-host and temporal sequence variation in a large CO1 sub-units from Anisakis simplex sensu stricto (Rudolphi, 1809) (Nematoda: Anisakisdae): implications for fisheries management. Mar Biol. 2007;151:695–702.
Ceballos-Mendiola G, Valero A, Polo-Vico R, Tejada M, Abattouy N, Karl H, et al. Genetic variability of Anisakis simplex s.s. parasitizing European hake (Merluccius merluccius) in the Little Sole Bank area in the Northeast Atlantic. Parasitol Res. 2010;107:1399–404.
Mladineo I, Poljak V. Ecology and genetic structure of zoonotic Anisakis spp. from Adriatic commercial fish species. Appl Environ Microbiol. 2014;80:1281–90.
Mattiucci S, Paggi L, Nascetti G, Abollo E, Webb SC, Pascual S, et al. Genetic divergence and reproductive isolation between Anisakis brevispiculata and Anisakis physeteris (Nematoda: Anisakidae). Int J Parasitol. 2001;31:9–14.
Mattiucci S, Nascetti G, Bullini L, Orecchia P, Paggi L. Genetic structure of Anisakis physeteris, and its differentiation from the Anisakis simplex complex (Ascaridida: Anisakidae). Parasitology. 1986;93:383–7.
Paggi L, Nascetti G, Webb SC, Mattiucci S, Cianchi R, Bullini L. A new species of Anisakis Dujardin, 1845 (Nematoda, Anisakidae) from beaked whales (Ziphiidae): allozyme and morphological evidence. Syst Parastitol. 1998;40:161–74.
Valentini A, Mattiucci S, Bondanelli P, Webb SC, Mignucci-Giannone AA, Colom-Llavina MM, Nascetti G. Genetic relationships among Anisakis species (Nematoda: Anisakidae) inferred from mitochondrial cox2 sequences, and comparison with allozyme data. J Parasitol. 2006;92:156–66.
Iglesias R, D’Amelio S, Ingrosso S, Farjallah S, Martínez-Cedeira JA, García-Estévez JM. Molecular and morphological evidence for the occurrence of Anisakis sp. A (Nematoda, Anisakidae) in the Blainville’s beaked whale Mesoplodon densirostris. J Helminthol. 2008;82:305–8.
Mattiucci S, Paoletti M, Webb SC. Anisakis nascettii n. sp. (Nematoda: Anisakidae) from beaked whales of the southern hemisphere: morphological description, genetic relationships between congeners and ecological data. Syst Parasitol. 2009;74:199–217.
Paggi L, Nascetti G, Cianchi R, Orecchia P, Mattiucci S, D’Amelio S, et al. Genetic evidence for three species within Pseudoterranova decipiens (Nematoda, Ascaridida, Ascaridoidea) in the north Atlantic and Norwegian and Barents seas. Int J Parasitol. 1991;21:195–212.
Vrijenhoek RC. Genetic differentiation among larval nematodes infecting fishes. J Parasitol. 1978;64:790–8.
Nascetti G, Cianchi R, Mattiucci S, D’Amelio S, Orecchia P, Paggi L, et al. Three sibling species within Contracaecum osculatum (Nematoda, Ascaridida, Ascaridoidea) from the Atlantic arctic-boreal region: reproductive isolation and host preferences. Int J Parasitol. 1993;23:105–20.
Orecchia P, Mattiucci S, D’Amelio S, Paggi L, Plötz J, Cianchi R, et al. Two new members in the Contracaecum osculatum complex (Nematoda, Ascaridoidea) from the Antarctic. Int J Parasitol. 1994;24:367–77.
Mattiucci S, Cipriani P, Paoletti M, Nardi V, Santoro M, Bellisario B, Nascetti G. Temporal stability of parasite distribution and genetic variability values of Contracaecum osculatum sp. D and C. osculatum sp. E (Nematoda: Anisakidae) from fish of the Ross Sea (Antarctica). Int J Parasitol Parasites Wildl. 2015;4:356–67.
Arduino P, Nascetti G, Cianchi R, Plötz J, Mattiucci S, D’Amelio S, et al. Isozyme variation and taxonomic rank of Contracaecum radiatum (v. Linstow, 1907) from the Antarctic Ocean (Nematoda, Ascaridoidea). Syst Parasitol. 1995;30:1–9.
Zhu X, D’Amelio S, Hu M, Paggi L, Gasser RB. Electrophoretic detection of population variation within Contracaecum ogmorhini (Nematoda: Ascaridoidea: Anisakidae). Electrophoresis. 2001;22:1930–4.
Mattiucci S, Cianchi R, Nascetti G, Paggi L, Sardella N, Timi J, et al. Genetic evidence for two sibling species within Contracaecum ogmorhini Johnston & Mawson, 1941 (Nematoda: Anisakidae) from otariid seals of Boreal and Austral regions. Syst Parasitol. 2003;54:13–23.
Li A-X, D’Amelio S, Paggi L, He F, Gasser RB, Lun Z-R, et al. Genetic evidence for the existence of sibling species within Contracaecum rudolphii (Hartwich, 1964) and the validity of Contracaecum septentrionale (Kreis, 1955) (Nematoda: Anisakidae). Parasitol Res. 2005;96:361–6.
Mattiucci S, Paoletti M, Olivero-Verbel J, Baldris R, Arroyo-Salgado B, Garbin L, et al. Contracaecum bioccai n. sp. from the brown pelican Pelecanus occidentalis (L.) in Colombia (Nematoda: Anisakidae): morphology, molecular evidence and its genetic relationship with congeners from fish-eating birds. Syst Parasitol. 2008;69:101–21.
RC is supported by a NERC studentship.
RC is supported by a NERC studentship.
Availability of data and materials
Ethics approval and consent to participate
Consent for publication
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Cole, R., Viney, M. The population genetics of parasitic nematodes of wild animals. Parasites Vectors 11, 590 (2018). https://doi.org/10.1186/s13071-018-3137-5
- Population genetics
- Population genomics
- Wild animals
- Population structure
- Parasite ecology