Min Hu1

Johan Höglund2

Neil B. Chilton1

Xingquan Zhu1, 3

Robin B. Gasser1

1Department of VeterinaryScience, The University ofMelbourne, Werribee,Victoria, Australia

2Department of Parasitology(SWEPAR), National VeterinaryInstitute and Swedish Universityof Agriculture Sciences,Uppsala, Sweden

3College of Veterinary Medicine,South China AgriculturalUniversity, Guangzhou,People’s Republic of China

Mutation scanning analysis of mitochondrialcytochrome c oxidase subunit 1 reveals limitedgene flow among bovine lungworm subpopulationsin Sweden

A mutation scanning approach was employed to investigate the population geneticstructure of the bovine lungworm, Dictyocaulus viviparus (Nematoda: Trichostrongyloi-dea), in southern Sweden. A total of 252 individual nematodes were collected fromcattle representing 17 farms. A portion of the mitochondrial cytochrome c oxidase sub-unit 1 gene (pcox1) was amplified from genomic DNA isolated from individual lung-worms by the polymerase chain reaction (PCR), and then subjected to single-strandconformation polymorphism (SSCP). Samples with distinct SSCP profiles were thensequenced. In total, 12 distinct pcox1 haplotypes (393 bp) were defined for the 252individuals, and pairwise sequence differences among the haplotypes ranged from0.3–2.3%. Average haplotype diversity and nucleotide diversity values were 0.16 and0.002, respectively. There was no particular correlation between pcox1 haplotypes andtheir geographical origin. The “overall fixation” indices FST and NST were calculated tobe 0.77 and 0.65, respectively. The results of this study revealed that both the mito-chondrial DNA sequence diversity within populations and the gene flow among popu-lations of D. viviparus were low. This is similar to findings for some parasitic nematodesof plants and insects, but distinctly different from gastrointestinal trichostrongyloidnematodes of domesticated ruminants considered to have relatively high levels ofgenetic diversity and gene flow. Such differences were interpreted to relate mainly todifferences in host movement as well as parasite biology, population sizes and trans-mission patterns, and should therefore be of epidemiological relevance.*

Keywords: Cytochrome c oxidase subunit 1 gene / Dictyocaulus viviparus (Nematoda) / Geneflow / Mitochondrial DNA / Mutation scanning / Population genetic structure EL 5120

1 Introduction

The bovine lungworm, Dictyocaulus viviparus (Strongy-lida), is a parasitic nematode causing severe, sometimesfatal bronchitis, particularly in calves. This disease (calledbovine husk or dictyocaulosis) is of major clinical impor-tance in many countries and causes substantial eco-nomic losses [1]. D. viviparus is dioecious and has a directlife cycle with a prepatent period of 3–4 weeks [2]. Theadult worms live in the bronchi, and the ovoviviparousfemales produce first-stage larvae (L1) which are shed inthe faeces of the host. Under optimal environmental con-ditions (i.e., temperature and humidity), it takes �5 days

for the L1s to develop to infective, third-stage larvae (L3).After ingestion by the host, the L3s invade the mesentericlymph nodes and are then transported as fourth-stage lar-vae (L4) via the lymph and blood to the lungs, where theyultimately develop to adults within �22 days.

Dictyocaulosis has major impact in temperate climaticzones with relatively high rainfall, ensuring the survival ofL3s in the environment. Traditionally, husk occurs incalves during the latter half of the first grazing season,and disease outbreaks occur most commonly in summerwhen the second generation of worms affects the host.After the end of patency (at the beginning of autumn),there is usually a rapid decrease in pasture contamination[3, 4]. While the vaccination of cattle with irradiated L3shas been effective in controlling bovine dictyocaulosisin mainland Europe, the demand for the vaccine has

Correspondence: Dr. Robin B. Gasser, Department of VeterinaryScience, The University of Melbourne, 250 Princes Highway,Werribee, Victoria 3030, AustraliaE-mail: robinbg@unimelb.edu.auFax: �61-3-97312366

Abbreviation: pcox1, portion of the cytochrome c oxidase sub-unit 1 gene

Electrophoresis 2002, 23, 3357–3363 3357

* Nucleotide sequences reported in this paper have been depos-ited in the DDBJ/EMBL/GeneBank databases under theaccession numbers AJ430568-AJ430579.

2002 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 0173-0835/02/2010–3357 $17.50�.50/0

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3358 M. Hu et al. Electrophoresis 2002, 23, 3357–3363

Table 1. Genetic diversity statistics for Dictyocaulus vivi-parus from cattle from 17 different farms in Swe-den (see Fig. 1).

FarmNo.

Haplotype code(Number of individuals)

Haplotypediversity (H)

Nucleotidediversity (�)

1 HP1 (26), HP12 (3) 0.19 0.00102 HP5 (9), HP7(1) 0.20 0.00263 HP3 (6), HP9 (2) 0.43 0.00784 HP3 (6), HP6 (3) 0.50 0.00265 HP1 (34) 0 06 HP1 (9) 0 07 HP2 (5) 0 08 HP1 (11) 0 09 HP3 (29), HP9 (9), HP10 (1) 0.40 0.0069

10 HP1 (2), HP8 (8) 0.36 0.006511 HP1 (6) 0 012 HP3 (7) 0 013 HP3 (25) 0 014 HP1 (4), HP9 (1) 0.40 0.006215 HP4 (8), HP10 (24), HP12 (2) 0.46 0.004416 HP11 (6) 0 017 HP5 (5) 0 0

FST = 0.77 NST = 0.65

The mitochondrial pcox1 haplotype codes, haplotypediversity (H) and nucleotide diversity (�) values as well asthe fixation indices, FST and NST, are listed.

recently decreased and, instead, it has been replaced bylong-acting anthelmintics which are used mainly in cattlein the first grazing season [5]. Consequently, young cattleare not adequately exposed to naturally occurring infec-tive larvae, resulting in reduced immunity in such animals.Also, this change in management has been associatedwith an increased susceptibility to disease in older cattle[6–9]. Other changes in husbandry practices, such as therecent emphasis on organic farming and associated pro-hibition of prophylactic treatment regimens [10], and atrend toward beef production in some parts of Europemay also contribute toward an increase in prevalence.

The unexpectedly high prevalence of dictyocaulosis inSweden, with �30–60% of organic farms studied havingat least one infected animal [11] has recently stimulatedinvestigations into the transmission patterns of D. vivi-parus, focused on designing improved control strategies.Central to the latter aspect is knowledge of the populationbiology and genetics of the parasite. For instance, knowl-edge of the population genetic structure and patterns ofgene flow allows predictions to be made about how aparasite population responds to changes in “natural,selective forces”, such as anthelmintic drugs, vaccinesand the adaptation of free-living larval stages to local cli-matic changes or differences.

The use of polymerase chain reaction (PCR) approaches,employing appropriate genetic markers, has significantlyfacilitated population genetic investigations [12–15].Mitochondrial DNA sequences are considered particular-ly useful for studying interspecific differences and intra-specific variation because of their high evolutionary ratesin animals and (proposed) maternal inheritance [16–19].For example, a recent study has shown the utility ofthe mitochondrial cytochrome c oxidase subunit 1 gene(cox1) for studying the structure of populations of hook-worms, employing a single-strand conformation poly-morphism (SSCP) approach [20]. Like hookworms, D. vivi-parus belongs to the order Strongylida, indicating theapplicability of the approach to the latter parasite.However, there are currently very limited sequence datafor the cox1 of D. viviparus (GenBank accession No.AF263474) and no published studies of its populationgenetic structure(s). Therefore, the aims of the presentstudy were to estimate, using an SSCP approach, the dis-tribution of sequence variation in a portion of the cox1among a large number of D. viviparus individuals fromdifferent farms in southern Sweden, in order to infer thepopulation genetic structure and gene flow for this parasite.

2 Materials and methods

2.1 Sampling, parasite collection and isolationof genomic DNA

Adults of D. viviparus were collected in abattoirs from thelungs of 1–3 cattle (1–49 nematodes per host individual)from each of 17 farms from the southern part of Sweden(Table 1; Fig. 1). The farms were separated by distancesranging from 4 to 670 km. Individual D. viviparus wereidentified by morphological and molecular approaches,as described recently [21]. Total genomic DNA wasextracted from a 1 cm portion of each individual worm,and purified using the QIAmp Tissue kit (Qiagen, SantaClarita, CA, USA), according to the protocol recom-mended. Each DNA sample was eluted into 400 �L of AEbuffer supplied in the kit and stored at –20�C.

2.2 Enzymatic amplification

A portion of the cytochrome c oxidase subunit 1 gene(pcox1) was amplified by PCR with 33P-endlabelledprimers JB3 (forward: 5’-TTTTTTGGGCATCCTGAGGTT-TAT-3’) and JB4.5 (reverse: 5’-TAAAGAAAGAACATAAT-GAAAATG-3’) [22]. The sequences of these primers havebeen verified to be sufficiently conserved among a rangeof nematodes (see [23]). PCR reactions (50 �L) were per-formed in 10 mM Tris-HCl, pH 8.4, 50 mM KCl, 4 mM

Electrophoresis 2002, 23, 3357–3363 Population genetic structure of bovine lungworm 3359

Table 2. Summary of nucleotide substitutions among the 12 mitochondrial pcox1 haplotypes (HP1 to HP12) of Dictyo-caulus viviparus in Sweden

Haplotype Distributionin the totalpopulation(in %)

Nucleotide position

6 3 5 6 1 1 1 1 1 2 2 2 2 2 26 1 6 1 4 5 7 9 0 3 6 7 8 8

4 4 9 1 8 1 7 2 2 5 8

HP1 36.5 A G A G G G G G A A G A A G GHP2 2.0 – – – – – – – – – T – – – – –HP3 29.0 G – – – – – – – – – – – – – –HP4 3.2 – – – – – – – – – – – – G – –HP5 5.6 – – – – – – – – G – – – – – –HP6 1.2 – – G – – – – – – – – – – – –HP7 0.4 – – G – A A – – – – – – – T –HP8 3.2 – A – A A – A A – – – G – – AHP9 4.8 – A – – A – A A – – – G – – AHP10 9.9 – A – – A – – – – – – – – – AHP11 2.4 – A – – A – – – – – A – – – AHP12 2.0 – – – – A – – – – – – – – T –

A dash indicates the same nucleotide as for haplotype HP1

Figure 1. Map of southern Sweden showing the geo-graphical location of the 17 farms, and the numbers andthe distribution of individual mitochondrial pcox1 haplo-types (HP1 to HP12) representing 252 Dictyocaulus vivi-parus individuals from cattle.

MgCl2, 250 �M each of dNTP, 50 pmol of each primerand 1 U Taq polymerase (Promega, Madison, WI, USA)in a thermocycler (Perkin Elmer Cetus, Norwalk, CT,USA) under the following conditions: 94�C for 5 min(initial denaturation), followed by 30 cycles of 94�C for30 s (denaturation), 55�C for 30 s (annealing), 72�C for30 s (extension), followed by a final extension at 72�Cfor 5 min. Samples without genomic DNA (no-DNA con-trols) or with bovine DNA (purified from musculature)were included in each amplification run, and in no casewere amplification products detected in these “nega-tive” controls. An aliquot (4 �L) of each PCR productwas examined by agarose gel electrophoresis (per-formed according to [24]) to establish amplification effi-ciency.

2.3 SSCP analysis

SSCP was used to screen the pcox1 amplicons forsequence variation among all individuals of each popula-tion. The SSCP method employed has the capacity todetect a single base difference for amplicons of �530 bp[25]. In brief, 10 �L of each amplicon were mixed with anequal volume of loading buffer (10 mM NaOH, 95% forma-mide, 0.05% bromophenol blue and 0.05% xylene cya-nole). After denaturation at 94�C for 2 min and subsequentsnap-cooling on a freeze-block (�20�C), 3 �L of eachsample were subjected to electrophoresis (7 W for 15 hat 18�C) in a 0.4 mm thick mutation detection enhance-ment gel matrix (FMC Bioproducts, Rockland, ME, USA).The conditions for electrophoresis were standardized for

3360 M. Hu et al. Electrophoresis 2002, 23, 3357–3363

optimal resolution of bands. After electrophoresis, gelswere dried on to blotting paper and subjected to auto-radiography for 24 h.

2.4 DNA sequencing and data analysis

Samples representing each of 17 farms and displayingvariable SSCP profiles were subjected to sequencing.Amplicons were purified over spin columns (Wizard

PCR-Prep; Promega) and subjected to automatedsequencing (BigDye chemistry, ABI). Sequencing wascarried out in both directions using primers JB3 andJB4.5 in separate reactions, according to the manufac-turer’s instructions. The program ARLEQUIN 2.0 [26] wasused to calculate the haplotype diversity (H) and nucleo-tide diversity (�). The F-statistics value (FST) was calcu-lated using the formula FST = (HT – HS)/HT, where HS is theaverage haplotype diversity in the populations, and HT isthe haplotype diversity for the population as a whole. NST,an analogue of FST at the DNA level, was calculated usingthe program Haplo 2, following Lynch and Crease [27]. A“minimum spanning tree” of the haplotypes was drawnusing ARLEQUIN 2.0. Amino acid sequences werededuced from nucleotide sequences using the programMacVector 4.1.4 employing the “invertebrate mitochon-drial genetic code” setting. Codon positions were deter-mined by comparative alignment with the cox1sequences of Caenorhabditis elegans, Ascaris suum [28],Ancylostoma duodenale and Necator americanus [23].Multiple lungworms (and their pcox1 sequences) fromthe same host were available for more than two hostsper site for some farms. In order to test whether lungwormsamples were distributed randomly, we used these sam-ples in an hierarchical F-statistics analysis on haplotypeidentity, and tested the significance of the variance com-ponent among hosts within a site using the ARLEQUIN2.0 program.

3 Results

No size variation was detectable on agarose gels amongthe pcox1 amplicons from all 252 individual adults of D.viviparus. Based on SSCP analysis of all individuals, 46samples representing the entire spectrum of profile varia-tion for all 17 farms were subjected to sequencing. Forthese samples, 12 distinct pcox1 sequence haplotypes(393 bp in length; 31% G�C) were defined (cf. Table 1).Alignment of all haplotypes revealed nucleotide variation(13 purine transitions and two transversions) at 15 align-ment positions (Table 2). Pairwise comparisons amongthe 12 haplotypes revealed sequence variation ranging

from 0.3–2.3%. Most of the nucleotide differences (n = 13;86.7%) were at the third codon position, whereas theremainder (n = 2; 13.3%) were at the first or second codonposition. Conceptual translation of individual pcox1 hap-lotypes revealed amino acid changes associated withsubstitutions at nucleotide positions 262, 272 and 285(cf. Table 2). For both haplotypes HP8 and HP9, theA�G transition at (first codon) position 262 with respectto HP1 resulted in a change from a methionine to a valine.For haplotype HP4, the same transition at (second codon)position 272 resulted in a change from an asparagine (inHP1) to a serine. For both haplotypes HP7 and HP12, theG �T transversion at (third codon) position 285 resultedin a change from a leucine to a phenylalanine.

The distribution of 12 pcox 1 sequence haplotypes isshown in Fig. 1. Haplotype HP1 occurred on 7 farms(Nos. 1, 5, 6, 8, 10, 11 and 14) in various parts of the studyarea, whereas haplotype HP3 occurred mainly on farms(Nos. 3, 4, 9, 12 and 13) in the southern and eastern partsthereof and on the island of Gotland (Fig. 1). HaplotypeHP9 occurred on three farms (Nos. 3, 9 and 14), haplo-types HP5, HP10 and HP12 were each present on twofarms and the other six haplotypes (HP2, HP4, HP6,HP7, HP8 and HP11) each occurred on one farm (Nos. 7,15, 4, 2, 10 and 16, respectively). Of the 17 farms, nine(Nos. 5, 6, 7, 8, 11, 12, 13, 16 and 17) had one haplotype,six (Nos. 1, 2, 3, 4, 10 and 14) had two haplotypes andtwo farms (Nos. 9 and 15) had three haplotypes. Therelationships among the 12 haplotypes and the numbersof individual haplotypes are displayed schematically ina minimum spanning tree (Fig. 2). While haplotypesHP1 and HP3 were found in 92 (36.5%) and 73 (29%) ofthe 252 D. viviparus individuals examined, the other tenhaplotypes were found in � 25 individuals (0.4–9.9%)(Fig. 2).

For selected samples, only 5.7% of the total variation inhaplotypic identity was distributed among hosts withinsites, and this value was not significantly different fromzero. This suggested that lungworm haplotypes were ran-domly distributed among hosts within farms. The statis-tics of genetic diversity of D. viviparus for the 17 farmsincluded in the study is shown in Table 1. Both haplotypediversity and nucleotide diversity within a farm were low,ranging from 0–0.5 and 0–0.0078, respectively, and aver-aging 0.16 and 0.002, respectively (Table 1). Taking all252 samples from all farms as one population, nucleotidediversity was calculated to be 0.006. Even though thegeographical distribution of the haplotypes did not indi-cate any particular pattern, FST and NST analyses revealedthat 77% of the haplotype diversity and 65% of the totalnucleotide sequence diversity were distributed amongpopulations (Table 1).

Electrophoresis 2002, 23, 3357–3363 Population genetic structure of bovine lungworm 3361

Figure 2. Relationships among the 12 mitochondrialpcox1 haplotypes (HP1 to HP12) representing Dictyocau-lus viviparus from cattle from 17 farms in Sweden. Thetotal number of samples representing each haplotype isindicated in brackets. The area of the circle is approxi-mately proportional to the number of samples. Hashmarks indicate nucleotide substitutions separating adja-cent haplotypes.

4 Discussion

According to the hypothesis of Price [29], the populationgenetic structures of parasites are expected to be asso-ciated with high levels of inbreeding, low intrapopulationgenetic variability and large genetic differentiation amongpopulations from different geographical regions or hosts.A number of studies of the population genetics of para-sitic nematodes either refute or support this hypothesisand indicate that parasitic nematodes usually show arange of overall genetic diversities and population geneticstructures [17, 18, 30–35].

Based on previous population genetic studies of parasiticnematodes, there are currently three types of structure.The first type is characterized by high genetic diversitieswithin populations and high gene flow among populations[17, 30, 35]. For example, in gastric trichostrongyloid

nematodes of ruminants, host movement is thought tocontribute substantially to the high gene flow amongpopulations, with large effective population sizes and ahigh evolutionary rate in mitochondrial DNA beingresponsible for high within-population diversities [17].The second type is characterized by highly structuredpopulations and low within-population diversity [31, 32,36]. An example of this is Heterorhabditis marelatus, aparasitic nematode of soil-dwelling insects, where smalleffective population sizes and restricted gene flow linkedwith its local transmission and ecology are thought to bethe explanations for the strict differentiation among popu-lations and low genetic diversity within populations andthe species as a whole [31]. The third structural type isexemplified by the population genetic structures of theascaridoid nematode of humans and pigs, Ascaris, andthe human hookworm, Necator americanus, which areboth characterized by significant genetic variation amongpopulations and no apparent correlation between geneticand geographical distance [18, 33, 34, 37]. Variable effec-tive population sizes of these parasites combined withhigh rates of host migration may explain this type of struc-ture (cf. [18]).

The present mutation scanning analysis revealed relati-vely low within-population variation in mitochondrialpcox1 and a relatively high degree of variation in geneticdiversity among subpopulations of D. viviparus in Swe-den, suggesting that the rates of gene flow among farmsare low. Interestingly, this genetic structure is similar tothose of the plant- and insect-parasitic nematodes(such as Meloidogyne arenavia and Heterorhabditismarelatus) but very distinct from related nematodes ofdomesticated cattle, such as Ostertagia ostertagi andHaemonchus placei, studied thus far [17, 30]. Consider-ing the differences in biology and epidemiology betweenD. viviparus and gastric trichostrongyloids of domesticruminants, there is a number of possible explanationsfor the population structure of the bovine lungworm inSweden.

It would appear that host movement does not play amajor role in the spread of D. viviparus between farms.Although cattle are transported between farms and themovement can sometimes be over vast distances, the“flow” of cattle differs depending on the kind of husban-dry system employed by producers. For example, individ-ual beef producers frequently recruit all of their calves onan annual basis, whereas dairy farmers usually producetheir own, although there may be limited exchange.Importantly, a relatively large proportion of farmers treatcattle with anthelmintics upon arrival to the farm, therebysignificantly reducing the risk of introducing lungworm ornew cohorts thereof. Also, young calves are introduced

3362 M. Hu et al. Electrophoresis 2002, 23, 3357–3363

prior to having been exposed to D. viviparus on pasture.Hence, host movement between or among farms may notcontribute significantly to the spread of the parasite.

Other epidemiological aspects would appear to be ofgreater significance. For example, the prevalence of lung-worm in Europe is usually lower than that of gastrointes-tinal nematodes [3, 38]. A recent epidemiological surveyof calves in Sweden revealed that Ostertagia was highlyprevalent (90–100% of cattle which have been on pas-ture), whereas the corresponding prevalence of Dictyo-caulus was �10% (for all age groups) upon stabling [11,39]. Also, the intensity of D. viviparus infection in cattle issignificantly lower [21] compared with gastrointestinalparasites where the intensity can be tens of thousands ofworms per host [40]. Although cattle of any age can beinfected with Dictyocaulus, the infection is most prevalentin yearlings, particularly towards the end of their first graz-ing season when it can be as high as �70% [3]. This ismainly related to the development of protective immunityprovoked by the infective larvae during their somaticmigration in the host. Consequently, the immunity againstD. viviparus is stronger compared with, for example,immunity against Ostertagia ostertagi (see [3]). Anotherbiological difference between the bovine lungworm andgastrointestinal nematodes relates to the survival of thelarval stages on pasture over winter. The “overwintering”of lungworm larvae on pastures in Europe is reported tobe relatively insignificant in parts of Europe (such as Den-mark, the Netherlands, Austria and Switzerland) [38], butit is relatively common in the UK and in Belgium [41]. InSweden, for instance, gastrointestinal nematodes survivewell on pastures over winter, whereas the consensus isthat lungworm larvae are poorer survivors. Furthermore,a recent study in Sweden demonstrated that wild rumi-nants, such as the roe deer and moose, are infected witha different species of Dictyocaulus [42], inferring thatneither of these two cervid hosts act as a reservoir of D.viviparus for the bovine host [21]. This is supported byexperimental evidence indicating that the cervid lung-worm, D. capreolus, is not transmissible to cattle [43].Taking all of this information into account, it is suggestedthat there are few opportunities in Sweden for differentsubpopulations (i.e., genetic variants) of D. viviparus tomove among farms, thus reflecting the low rate of geneflow calculated. Consequently, the findings suggest thatthe parasite may entertain a high level of inbreeding withina population, which would also explain the relatively lowgenetic diversity within subpopulations (i.e., farms).

In conclusion, this study reveals that the populationgenetic structure of D. viviparus in Sweden is more similarto some parasitic nematodes of plants and insects [31,32] than to related gastric nematodes (of the same order

and superfamily) of domesticated ruminants, which canbe explained by significant biological and ecological dif-ferences. The results of this study should have thereforeimportant epidemiological implications. Whether the pop-ulation genetic structure of D. viviparus in Sweden is thesame as that in other countries is currently unknown, andis thus worthy of detailed investigation employing the mo-lecular tools used herein.

We are grateful to M. Blouin for discussions and com-ments, and to P. Brown for forwarding the Haplo 2 pro-gram to us. Thanks also to J. Wood for assistance withsequencing. Funding support was provided to J.H. bythe Swedish Farmer’s Foundation for AgriculturalResearch and the Swedish Meats Association and FOR-MAS, and to R.B.G. through the Australian ResearchCouncil and other sources. M.H. has been a recipient ofa postgraduate scholarship from the Faculty of VeterinaryScience of The University of Melbourne.

Received June 15, 2002

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