Molecular phylogenetic analysis of the genus Strongyloides and related nematodes

International Journal for Parasitology 32 (2002) 1507–1517 www.parasitology-online.com

Molecular phylogenetic analysis of the genus Strongyloides and related nematodes q Mark Dorris a, Mark E. Viney b, Mark L. Blaxter c,* a

Centre for Marine Biodiversity and Biotechnology, Heriot–Watt University, Riccarton, Edinburgh EH14 4AS, UK b School of Biological Sciences, University of Bristol, Woodland Road, Bristol BS8 1UG, UK c Ashworth Laboratories, Institute of Cell, Animal and Population Biology, King’s Buildings, University of Edinburgh, Edinburgh EH9 3JT, UK Received 13 May 2002; received in revised form 23 July 2002; accepted 23 July 2002

Abstract Strongyloides spp., parasitic nematodes of humans and many other terrestrial vertebrates, display an unusual heterogonic lifecycle involving alternating parasitic and free-living adult reproductive stages. A number of other genera have similar lifecycles, but their relationships to Strongyloides have not been clarified. We have inferred a phylogeny of 12 species of Strongyloides, Parastrongyloides, Rhabdias and Rhabditophanes using small subunit ribosomal RNA gene (SSU rDNA) sequences. The lineage leading to Strongyloides appears to have arisen within parasites of terrestrial invertebrates. Inferred lifecycle evolution was particularly dynamic within these nematodes. Importantly, the free-living Rhabditophanes sp. KR3021 is placed within a clade of parasitic taxa, suggesting that this species may represent a reversion to a non-parasitic lifecycle. Species within the genus Strongyloides are very closely related, despite the disparity of host species parasitised. The highly pathogenic human parasite Strongyloides fuelleborni kelleyi is not supported as a subspecies of the primate parasite S. fuelleborni fuelleborni, but is most likely derived from a local zoonotic source. q 2002 Australian Society for Parasitology Inc. Published by Elsevier Science Ltd. All rights reserved. Keywords: Strongyloides; Parastrongyloides; Rhabdias; Rhabditophanes ribosomal RNA; Molecular phylogeny

1. Introduction The phylum Nematoda has a large number of parasitic groups, and molecular phylogenetic analysis suggests that there have been multiple independent events of gain of parasitism of both animals and plants (Blaxter et al., 1998, Dorris et al., 1999). Importantly, it has been possible to identify free-living sister taxa for most (but not all) of the major parasitic nematode taxa. A comparison of the evolutionary origins of different parasitic groups may reveal common patterns that will illuminate prerequisites for parasitism, and the dynamics of its acquisition. The genus Strongyloides comprises over 40 species of nematodes parasitic in vertebrates (Grove, 1989; Viney, 1988). While a few species with amphibian, reptile, and bird hosts are known, the vast majority parasitise mammals, q Sequences reported in this paper have been deposited in the EMBL/ GenBank/DDBJ databases under the accession numbers AJ417021– AJ417032. The alignments used have been deposited in Treebase and EMBL (Accessions ALIGN_000446, ALIGN_000467, and SN795). * Corresponding author. Tel.: 144-131-650-6760; fax: 144-131-6507489. E-mail address: [email protected] (M.L. Blaxter).

many of which are domesticated, including cats, dogs and livestock. The wide range of vertebrate hosts parasitised by Strongyloides species could have arisen through either ancient parasitism and subsequent co-speciation with the host, and/or evolution of parasitism followed by horizontal transfer between hosts. Three species infect humans. Strongyloides stercoralis is a human pathogen with a worldwide distribution; over 600 million infections are estimated worldwide (Chan et al., 1994). Strongyloides fuelleborni fuelleborni infects non-human primates in Africa, but can also infect humans (Ashford and Barnish, 1989). A third, potentially fatal human strongyloidiasis was discovered in New Guinea, attributed to the subspecies Strongyloides fuelleborni kelleyi (see Kelly et al., 1976). Strongyloides fuelleborni kelleyi was distinguished from S. f. fuelleborni by the appearance of the peri-vulval cuticle of the parasitic females, the position of the phasmidial pore of free-living males and by the geographical isolation of S. f. kelleyi (see Viney et al., 1991). Development of treatments and increased understanding of the biology of important parasitic species can be approached by the study of model species. The rodent parasite Strongyloides ratti is thought to provide such a model

0020-7519/02/$20.00 q 2002 Australian Society for Parasitology Inc. Published by Elsevier Science Ltd. All rights reserved. PII: S 0020-751 9(02)00156-X

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for human strongyloidiasis, but it is not known how closely related this species is to the human pathogens. On a larger scale, a relatively close relationship has been shown between Strongyloides and the insect pathogenic nematode genus Steinernema, based on molecular (Blaxter et al., 1998; Dorris et al., 1999) and morphological (De Ley, 1995) evidence. These two genera were classified within Rhabditina due to the occurrence of ‘rhabditiform’ larvae, but recently have been grouped with Rhabdiasidae (including anuran lung parasites) and Alloionematidae (free-living microbivores, and arthropod and gastropod parasites) to form a new superfamily Strongyloidoidea (De Ley, 1995; De Ley and Blaxter, 2001). The interrelationships among species within this proposed superfamily, and the relationships of the superfamily to other nematodes, including freeliving taxa, are unresolved. Species within the Strongyloidoidea have direct, but often complex, lifecycles (Fig. 1) (Harvey and Viney, 2001; Viney, 1996). While parasitic stages of Strongyloides are all female, and reproduce parthenogenetically (Viney, 1994), in Parastrongyloides, parasitic males are found and in Rhabdias free-living hermaphrodites and a parasitic sexual phase are found (Anderson, 2000). In Strongyloides, entry into the parasitic phase is obligatory but some species can undergo more than one free-living cycle in culture (Yamada et al., 1991). Parastrongyloides trichosuri (from Australian common possums) can have

many free-living sexual cycles with careful maintenance of culture conditions (W. Grant, personal communication), while Rhabdias bufonis (a lung parasite of European frogs) has a single, hermaphroditic free-living cycle (Smyth and Smyth, 1980; Spieler and Schierenberg, 1995). Strongyloides has attracted much interest because the developmental choice between lifecycles may represent an intermediate between a free living and wholly parasitic phenotype (Viney, 1999), but this lifecycle with alternating generations must be currently selectively advantageous, and maintained for reasons of fitness (Read and Skorping, 1995). The patterns of lifecycle history displayed by different species are adaptations to particular host and environment combinations, but may also be constrained by their phylogenetic history. The evolution of the parasitic phenotype requires coordinate acquisition of many novel traits and thus reversion to a free-living state is thought to be unlikely. Reversion has been proposed for flagellated protists within the Diplomonadida (Siddall et al., 1992) but members of the phylum Nematoda with alternating life histories provide likely candidates for parasitic reversal within the Metazoa (Poulin, 1998). In order to provide a framework from which to address some of these issues we present a molecular phylogenetic analysis of the genus Strongyloides and close relatives based on small subunit ribosomal gene (SSU rDNA) sequences.

Fig. 1. Lifecycle variation in Strongyloides and related nematodes. A cartoon depicting the diversity of alternating lifecycles of nematodes related to Strongyloides spp. The taxa are linked by a tree derived from the results of this study (but note that Strongyloides planiceps was not sampled). L3i, infective third stage larvae. Steinernema species are bacteriovores that infect an insect ‘host’ with a pathogenic bacterium on which they feed. The anuran parasite Rhabdias has alternating free living and parasitic generations. Rhabditophanes sp. is grown in the laboratory as a free-living bacteriovore but may have a paratenic arthropod associate in the wild. Parastrongyloides can undergo multiple free-living cycles in addition to the parasitic one, but only in carefully controlled culture conditions. While Strongyloides ratti and Strongyloides stercoralis have but one free living cycle, S. planiceps has been reported to be able to undergo multiple free-living cycles in culture.

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2. Materials and methods 2.1. Source of nematode materials Taxa sampled for phylogenetic analysis are described in Table 1. Fresh, frozen S. stercoralis infective stage larvae (L3i) were supplied by Prof G. Schad (School of Veterinary Medicine, University of Pennsylvania, PA, USA) and the complete SSU rDNA gene was amplified and sequenced. A previously published S. stercoralis sequence was a hybrid containing contaminating fungal sequence (Putland et al., 1993), and thus this species was resequenced (Dorris and Blaxter, 2000). Formalin-fixed specimens of Strongyloides cebus, S. fuelleborni fuelleborni, S. fuelleborni kelleyi, Strongyloides venezuelensis and a Strongyloides sp. from spitting cobra were sampled. The cobra-derived Strongyloides was not identified with species as only infective larvae were available. There are two described snake Strongyloides, Strongyloides serpentis and Strongyloides gulae, from the same host, the water snake Natrix cyclopion which differ only in the size of the adults. The cobra-derived infective larvae were collected from the Liverpool School of Tropical Medicine herpetarium and thus cross-infection from other snake hosts cannot be ruled out. DNA extracted from bulk formalin-fixed specimens of

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Strongyloides westeri, Strongyloides suis and Strongyloides papillosus were supplied by Dr T. Moore and Dr T. Nutman (Laboratory for Parasitic Diseases, NIH, Bethesda, MD, USA). Parastrongyloides trichosuri genomic DNA was supplied by Dr W. Grant (Flinders University, South Australia). Steinernema kari genomic DNA was supplied by Dr A. Reid (CABI, St. Albans, UK). A culture of Rhabditophanes sp. KR3021, originally isolated by Prof A. Rose near Vancouver, British Columbia, Canada, was supplied by Dr W.K. Thomas (University of Missouri, KS, USA). The same strain was sequenced by Felix et al. (2000). Live and ethanol-fixed R. bufonis were supplied by Prof E. Schierenberg (Zoological Institute, Ko¨ ln, Germany). Caenorhabditis elegans N2 was obtained from the Caenorhabditis Genetics Center (St. Paul, MN, USA) 2.2. Development of a method for extraction and amplification of DNA from formalin-fixed samples Optimisation of DNA extraction and amplification was essential to permit analysis of small numbers of formalinfixed specimens (Herniou and Pearce, 1998). PCRs were set up using dedicated pipettes in a PCR-dedicated, UV-sterilisable flow hood. Downstream manipulations (gel electrophoresis, cloning, sequencing) were carried out in a

Table 1 Taxa sampled a Taxon(species name and strain identification code)

Source material

Host

Length of SSU rDNA analysed

GenBank accession number

Strongyloides stercoralis Strongyloides cebus Strongyloides fuelleborni kelleyi Strongyloides fuelleborni fuelleborni Strongyloides venezuelensis Strongyloides sp. ex snake Strongyloides westeri Strongyloides suis Strongyloides papillosus Strongyloides ratti Parastrongyloides trichosuri Rhabdias bufonis Rhabditophanes sp. KR3021 Steinernema kari Steinernema carpocapsae Acrobeles complexus WUCG2 Acrobeles sp. PS1156 Acrobeloides bodenheimeri PS2160 Cephalobus oryzae PS1165 Halicephalobus gingivalis PDL0017 Panagrellus redivivus PS1163 Panagrolaimus sp. PS1159 Panagrobelus stammeri PDL0024 Cervidellua alutus PDL0004 Zeldia punctata Ascaris lumbricoides Brumptaemilius justini Gnathostoma neoprocyonis

Frozen material Fixed material Fixed material Fixed material Fixed material Fixed material DNA from fixed material DNA from fixed material DNA from fixed material GenBank Fresh material Fresh material Fresh material Fresh material GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank GenBank

Dog laboratory host South American non-human primate Papua New Guinea human African primate Rat Spitting cobra Horse Pig Rabbit Rat Australian common possum European common toad Free-living Insect pathogen Insect pathogen Free-living Free-living Free-living Free-living Free-living Free-living Free-living Free-living Free-living Free-living Human Millipede Seal

1,200 330 330 332 329 332 330 330 330 1,200 1,153 1,197 1,200 1,228 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200 1,200

AF279916 AJ417025* AJ417029* AJ417030* AJ417026* AJ417031* AJ417032* AJ417028* AJ417027* AF036605 AJ417024* AJ417022* AF202151 AJ417021* AF036604 U81577 U81576 AF202162 AF034390 AF202156 AF083007 U81579 AF202153 AF202152 U61760 U94366 AF036589 Z96947

a

*Sequences determined in this study.

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Table 2 SSU rDNA primers used for PCR and sequencing a Name G18S4 SSUA SSU22F SSU22R SSU9F SSU9R SSU24F SSU26R SSU23F SSU23R SSU13R SSU18P SSUDR a

Usage PCR and sequencing PCR and sequencing Sequencing PCR and Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing Sequencing PCR and sequencing PCR and sequencing

Position*

Sequence

0

5 end 39–57 411–428 429–411 573–591 584–565 868–887 927–907 1,280–1,298 1,298–1,280 1,438–1,419 3 0 end 1,213–1,194**

GCTTGTCTCAAAGATTAAGCC AAAGATTAAGCCATGCATG TCCAAGGAAGGCAGCAGGC GCCTGCTGCCTTCCTTGGA CGCGGTAATTCCAGCTCCA AGCTGGAATTACCGCGGCTG AGRGGTGAAATYCGTGGACC CATTCTTGGCAAATGCTTTGC ATTCCGATAACGAGCGAGA TCTCGCTCGTTATCGGAAT GGGCATCACAGACCTGTTA TGATCCWKCYGCAGGTTCAC CATAAAAGTCTCGCTCGTTA

*Numbering as for C. elegans SSU rDNA sequence except ** from S. ratti sequence.

separate laboratory. Caenorhabditis elegans adult hermaphrodites were fixed in 4% formalin and subsequently dehydrated in glycerol, mimicking the fixation procedure for slide preparation. After 14 days in glycerol, nematodes were rehydrated and incubated overnight in GTES buffer (100 mM glycine, 0.05% SDS, 10 mM TrisHCl, 1 mM (EDTA)). Nematodes were then washed in 10 mM TrisHCl pH 8.0. Individual nematodes were subjected to digestion and direct PCR. Different primer sets were used to determine the maximum size of product that could be amplified (Table 2). Nematodes fixed for long periods, such as specimens from slides (from the study of Viney, 1988), were still structurally intact after 24 h incubation with proteinase K. To extract PCR-accessible DNA from these specimens, nematodes were picked from a slide (on which they were mounted in glycerol after formalin fixation 10 years back). They were rehydrated in reducing concentrations of ethanol (100–75–50–25–0%) and incubated at room temperature overnight in GTES buffer. The specimens were then freeze-fractured in liquid N2 and ground with a micro mortar and pestle. Genomic DNA was extracted for subsequent PCR using the Nucleon HT extraction method following the manufacturer’s recommended protocols (Scotlab). This method differs from others in that the resin used binds (and removes) protein and other impurities rather than binding DNA. 2.3. DNA sequencing SSU rDNA gene fragments (1,500–1,700 bp) were directly amplified by PCR from fresh specimens of individually identified nematodes after protease digestion, or from nematode DNA samples, using universal SSU primers (Table 2). Only 1,229 bp of SSU sequence was amplified from R. bufonis. Three hundred and thirty base pairs from the 5 0 end of the SSU gene was amplified from formalinfixed Strongyloides (Table 1). PCR products from fresh

material were purified from agarose gels using a glass matrix method (Hybaid) and sequenced directly using fluorescent dye terminators (Perkin–Elmer corporation) in both directions using internal conserved primers (Table 2). 5 0 partial fragments amplified from formalin-fixed samples were cloned into pMOSblue (Amersham). At least three individual clones were sequenced in both directions using the SSUA or G18S4 forward primers and SSU22R reverse primer. A majority rule consensus was derived for each fragment. 2.4. Alignment and phylogenetic analyses Two separate datasets were compiled from partial SSU rDNA sequences. All sequences used for phylogenetic analysis were added to the alignment of Blaxter et al. (1998), which contains SSU rDNA sequence from taxa sampled across the phylum Nematoda. The first data set comprised ,75% of the full length SSU rDNA sequence, limited by 1,229 bp of sequence (1,270 characters aligned including gaps) from R. bufonis. Within this data set, Strongyloidoidea was represented by two Strongyloides spp., one Parastrongyloides, one Rhabditophanes and two Steinernema spp. Rhabdias bufonis was also included as were four Panagrolaimidae and six Cephalobidae for comparison. One representative each from Ascaridida, Spirurida and Rhigonematida was included as outgroups. The second data set comprised 380 characters including gaps and incorporates 5 0 partial sequence from additional Strongyloides spp. In addition to 10 Strongyloidess sequences, P. trichosuri, Rhabditophanes sp. KR3021, R. bufonis and two steinernematids were included. Phylogenetic analyses were carried out using PAUP* 4.0 beta 10 (Rogers and Swofford, 1999; Swofford, 1993, 1999; Swofford et al., 1996) and MRBAYES (Huelsenbeck and Ronquist, 2001; Huelsenbeck et al., 2001). The strategy for phylogenetic analysis, using maximum parsimony, neighbour joining and maximum likelihood methods in PAUP*

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followed that described by Rogers and Swofford (1999). For maximum likelihood and neighbour joining analyses, ModelTest (v3.06) (Posada and Crandall, 1998) suggested that for both datasets the GTR 1 I 1 gamma (general time reversible plus proportion of invariant sites plus gamma distribution of rate parameters) model best described the data. Support for the tree topologies was assessed by 1,000 bootstrap replicates. Bayesian Markov chain Monte Carlo analysis was carried out using MRBAYES (v2.01) using four chains with six nucleotide substitution rates, a gamma distribution of rate variation between sites, and 1,000,000 replicates. Chain states and tree topology were sampled every 100 iterations. The first 500 saved states were discarded before consensus trees were computed.

3. Results 3.1. Development of a method for DNA extraction for PCR from formalin-fixed nematodes Methods for extraction of DNA for PCR were tested on formalin-fixed C. elegans. Standard methods for DNA extraction from nematodes were not effective in releasing DNA accessible for PCR amplification from formalin-fixed specimens, while single nematodes yielded amplification products in the absence of fixation. For nematodes fixed briefly (see Materials and methods), incubation in GTES buffer released DNA that was accessible for PCR amplification (Fig. 2). Products of #1,000 bp were readily obtained

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from recently fixed and dehydrated specimens but full length SSU PCR products (1,700 bp) were not obtained. Direct sequencing of these PCR products yielded readable sequence corresponding to the C. elegans SSU gene. Bulk DNA samples prepared from formalin-stored nematodes were not accessible to PCR without further purification: the longest products obtained were ,150 bp. Strongyloides suis genomic DNA prepared from bulk fixed nematodes was diluted 1:10 with GTES, incubated overnight, subjected to Nucleon HT extraction, and the primer set SSUA/SSU22R (Table 2) used to amplify ,400 bp of the 5 0 end of the SSU (Fig. 2B). A high yield of product was obtained, but multiple molecular species were amplified. Direct sequencing of PCR products yielded poor or no sequence (see Herniou and Pearce, 1998). Cloning of PCR products permitted sequencing of individual molecular species, which derived from the expected 5 0 region of the SSU. PCR products from samples that had been exposed to formalin for long periods were therefore cloned before sequencing. Long-term formalin fixation of tissues appears to detrimentally affect DNA such that it is only poorly available for amplification, and the DNA that is amplifiable is heterogeneous in sequence. It is likely that covalent modifications to the nucleic acid, or induced errors in polymerase extension, result in the populations of amplicons that differ by base deletions, making sequencing of PCR amplicon mixtures uninformative. For nematodes fixed on slides, 1–20 individual nematodes were selected by hand from the slide, and processed

Fig. 2. PCR amplification of small subunit ribosomal RNA gene fragments from formalin-fixed nematodes. (A) PCR from Caenorhabditis elegans fixed in formalin. Primer sets used were as follows (see Table 2): lane 1: SSUA/SSU26R, lane 2: SSU22F/SSUDR, lane 3: SSU9F/SSU18P, lane 4: SSUA/SSU26R (positive control; amplification from fresh C. elegans), lane 5: SSU22F/SSU26R, lane 6: SSUA/SSU22R, lane 7: SSUA/SSU9R, and lane 8: SSU9F/SSUDR. The PCR product in lane 3 was sequenced using the primer SSU9F; no differences from the published C. elegans SSU rDNA sequence were observed (data not shown). Primer sequences are shown in Table 2. The sizes of molecular markers (lanes m) are shown. (B) PCR products from fixed specimens from slides. Primer sets used were as follows (see Table 2): lanes 1–3: SSUA/SSU22R; lanes 4–6: SSUA/SSU18P. Lanes 1 and 4 contain products amplified from formalinfixed Strongyloides fuelleborni fuelleborni samples and lanes 2 and 5 contain products amplified from Strongyloides suis genomic DNA. Starting material from both sources was incubated in GTES (see text) and genomic DNA was extracted using the Nucleon HT extraction kit. Lanes 3 and 6 are positive controls and contain products from unfixed C. elegans. Very faint bands were visible in lanes 4 and 5.

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as above. For each PCR from fixed material that was positive (and where the concurrent negative control was negative), 96 individual clones were selected and placed on a microtitre plate. These were screened by PCR using specific primers directed to known contaminants. From the remaining clones, three were picked for sequencing. If these three did not yield a single consensus, additional clones were sequenced. Cloning and PCR screening of products proved essential for these samples, as many of the clones had fungal, tree (the wooden toothpick fragments used to support the coverslip) or human (operator, or original handling contamination) origins. The problem of amplification of non-nematode DNA had previously led to chimaeric sequences being analysed (Putland et al., 1993). Ten Strongyloides species were sequenced as well as one Rhabdias, one Parastrongyloides and one Steinernema. Rhabditophanes sp. KR3021 was also processed, and the sequence was found to be identical to that obtained by others (Felix et al., 2000). 3.2. The relationship of Strongyloides to other nematodes with alternating lifecycles Analysis of the first dataset of near-full length SSU sequences from taxa representing Strongyloides, Parastrongyloides, Rhabdias, Steinernema, Rhabditophanes and a selection of cephalobid outgroups using maximum parsimony and neighbour joining methods yielded identical trees with high bootstrap support (Fig. 3). No more likely trees were found in a maximum likelihood search. Bayesian Markov chain Monte Carlo analysis yielded a topology identical to the other methods, except that the node distinguishing (Steinernema (Rhabdias (Strongyloides, Parastrongyloides, Rhabditophanes))) from ((Rhabdias, Steinernema) (Strongyloides, Parastrongyloides, Rhabditophanes)) was not strongly supported. Examination of the maximum likelihood models being searched by the Bayesian Markov chain Monte Carlo analysis showed that they robustly (and repeatedly) converged on a very different set of maximum likelihood parameters from that determined heuristically. For example, the mean estimated gamma parameter for the last 100,000 generations of Bayesian Markov chain Monte Carlo analysis was 0.429, compared to 0.902 estimated by the method of Rogers and Swofford (1999). This difference in model parameters probably underlies the difference in the trees supported. However, all methods converge on very similar trees with otherwise comparable levels of support, and this is strongly suggestive of a robust topology. The tree produced by maximum likelihood is shown in Fig. 3A and a cladogram showing levels of support by neighbour joining bootstrap employing maximum likelihood parameters based on the maximum likelihood tree is shown in Fig. 3B. The topology found by Bayesian Markov chain Monte Carlo analysis is shown with support levels in Fig. 3C. The molecular analysis supports the proposal that (Steinernema 1 Rhabdias 1 Rhabditophanes 1

Fig. 3. The phylogenetic relationships of the Strongyloidoidea. (A) Phylogram derived from a maximum likelihood analysis of aligned SSU rDNA sequences. (B) Cladogram of the tree in (A), showing support levels derived from 1,000 bootstrap resamplings analysed using neighbour joining with maximum likelihood parameters. The numbers above each branch show bootstrap support. Branching orders supported at less than 50% are collapsed to form polytomies. (C) Cladogram of the consensus of 9,500 sampled trees from a 1,000,000 generation Bayesian Markov chain Monte Carlo analysis of the same dataset. Posterior probability values (as %) are given above supported nodes.

Parastrongyloides 1 Strongyloides) are members of a single higher taxon, Strongyloidoidea, rather than disparate members of Rhabditina and Cephalobina. The Strongyloidoidea are robustly placed within Clade IV of Blaxter et al. (1998). This concurs with a taxonomically more limited analysis, which included the erroneous S. stercoralis sequence and Rhabditophanes (Felix et al., 2000). Panagrolaimidae were found to be a sister clade to Strongyloidoidea, in contrast to a previous analysis in which Panagrolaimidae were closer to Strongyloides with Steinernematidae as a sister group. Within Strongyloidoidea, the branching order was found to be (Steinernema (Rhabdias (Rhabditophanes

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(Parastrongyloides, Strongyloides)))) in maximum parsimony, neighbour joining, heuristic search maximum likelihood and Bayesian Markov chain Monte Carlo analyses. Branch lengths within Cephalobidae are uniformly shorter than those within the Panagrolaimidae and Strongyloidoidea. 3.3. The interrelationships of Strongyloides species Ten species of Strongyloides were sampled from a representative wide host range, including a snake, bovid, rodents, primates and all three recognised parasites of humans (Table 1). The Strongyloides SSU sequences were all very similar which made the resolution of their phylogeny problematic with distance and likelihood methods: many branch lengths are inferred to be very short. Indeed, specimens of S. cebus, S. papillosus and S. f. kelleyi yielded the same sequence, as did S. stercoralis, S. f. fuelleborni and Strongyloides sp. ex snake. These sequences are unlikely to be laboratory contamination artefacts as the PCRs were performed independently, at different times. Bayesian Markov chain Monte Carlo, maximum parsimony and neighbour joining methods converged on the same tree with comparable levels of support. The maximum likelihood tree is shown in Fig. 4A and a cladogram showing levels of support by neighbour joining bootstrap employing maximum likelihood parameters based on the maximum likelihood tree is shown in Fig. 4B. The Bayesian Markov chain Monte Carlo tree is shown in Fig. 4C. The genus can be split into two clades: one containing Strongyloides sp. ex snake, S. stercoralis and S. f. fuelleborni, and one containing S. ratti, S. suis, S. venezuelensis, S. cebus, S. f. kelleyi and S. papillosus. Strongyloides westeri arises from a basal polytomy. 3.4. A molecular synapomorphy for two clades of Strongyloides Closer analysis of the SSU sequences from the Strongyloides species examined here identified putative synapomorphies within the E9-2 hairpin of the V2 variable region of the SSU sequence (Fig. 5). Reanalysis of the alignment excluding this short region still returns the two clades (data not shown). 4. Discussion 4.1. Extraction of DNA from formalin-fixed specimens Formalin fixation chemically alters DNA, and makes PCR amplification of target genes difficult (Herniou and Pearce, 1998). In particular, any non-fixed DNA contaminating an extraction from otherwise formalin-fixed material will be preferentially amplified. This contamination could arise during storage, or during manipulation for PCR. For nematode material recently fixed in formalin, we were able

Fig. 4. The phylogenetic relationships within the Strongyloidoidea. (A) Phylogram derived from a maximum likelihood analysis of aligned SSU rDNA sequences. (B) Cladogram of the tree in (A), showing support levels derived from 1,000 bootstrap resamplings analysed using neighbour joining with maximum likelihood parameters. The numbers above each branch show bootstrap support percentages. Branching orders supported at less than 50% are collapsed to form polytomies. (C) Cladogram of the consensus of 9,500 sampled trees from a 1,000,000 generation Bayesian Markov chain Monte Carlo analysis of the same dataset. Posterior probability values (as %) are given above supported nodes.

to recover DNA amplicons up to 1,000 bp. However, for Strongyloides specimens that were stored for over 10 years, this method in itself was insufficient. Nematodes fixed for long periods were refractory to protease digestion as they were still structurally intact after 24 h incubation with proteinase K. Formaldehyde fixation of the nematode cuticle may be irreversible even in the presence of the aggressive detergent SDS. A methodology involving careful extraction of clean DNA, attention to very clean PCR conditions and cloning and screening of resultant amplified fragments allowed us to derive sequences for the fixed material.

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mic DNA and informative sequence from long-fixed specimens is achievable.

4.2. Composition and the origins of parasitism in Strongyloidoidea

Fig. 5. A molecular synapomorphy supporting two clades within Strongyloides. (A) A cartoon of the E9 region of the SSU rRNA molecule from Strongyloides ratti showing proposed secondary structures. The E9-2 stemloop is boxed. (B). The E9-2 stem-loop section of the SSU rDNA alignment showing the proposed synapomorphy for the ‘stercoralis’ (outlined) and ‘cebus’ (grey shading) clades.

Sequencing of multiple clones yielded consistent sequence, with individual clones differing in only one or two bases .300. This level of difference should not affect the conclusions we have drawn, as the differences observed between species groups within Strongyloides were in excess of this. The quality and history of the extracted DNA is paramount in this process, as previous authors have found differences of up to 10% in sequence from formalin-fixed specimens of the same species (Littlewood, 1999, Nadler, 1999). Our data were much less variable, perhaps because we optimised extraction of very clean DNA, rather than maximising DNA yield. Potential sources of cross-contamination noted previously (Herniou and Pearce, 1998) can be ruled out in our data set, as there was no potential for mixing of sample storage medium. It is also possible that formalin fixation could have a non-random effect on DNA, causing repeatable changes in subsequently determined sequence. This was not evident from our trials with C. elegans but remains a testable possibility for long-term fixed material. Although not a trivial procedure, extraction of quality geno-

Strongyloides spp. were traditionally classified as rhabditids, placed within the Rhabditina or Rhabditoidea. Previous molecular analyses supported cephalobid origins for Strongyloides, placing them within a clade (‘Clade IV’) that included cephalobids, aphelenchs, tylenchs and panagrolaims (Blaxter et al., 1998, 2000; Felix et al., 2000). The proposal for a superfamily (Strongyloidoidea) comprising these taxa (De Ley, 1995; De Ley and Blaxter, 2001) is strongly supported by the molecular evidence presented here in which the genera Strongyloides and Steinernema, the family Strongyloididae, and superfamily Strongyloidoidea are all strongly supported monophylies. Rhabditophanes is shown to be monophyletic with Strongyloididae. While insect associations have been described for members of Strongyloidoidea, an insect-parasitic state preceding vertebrate parasitism is unlikely as there are no true insect parasites within the group. Steinernematids kill their hosts via a toxic bacterial symbiont and their association is thus with dead insects. The free-living stages of the vertebrate parasites are not known to associate with insects, phoretically or otherwise, and infection of the host is almost exclusively by a percutaneous route, suggesting that an ancestral oral transmission by ingestion of an infected arthropod is unlikely. Rhabditophanes larvae have a phoretic association with beetles, but only by external attachment (Rhu¨ m, 1956). They are not found associated with the internal body cavity. Some members of Alloionematidae are snail parasites with alternating lifecycles (Cabaret and Morand, 1990) and Rhabdias parasites use snails as transport hosts (Smyth and Smyth, 1980). It is possible that the first parasites in this group had gastropod hosts. Thus, the origins of vertebrate parasitism in Strongyloididae may lie with the ability to recognise and attach firmly to a host. Comparative neuroanatomical analysis suggests a possible mechanism for the evolution of percutaneous transmission observed in Strongyloides. In C. elegans, the sensory endings of the cephalic neurons are exposed frontally (Herman, 1996) but the homologous sensory endings in Strongyloides have a more radial exposure (Ashton et al., 1995, 1999; Fine et al., 1997). C. elegans exhibits nosetouch avoidance (Herman, 1996; Kaplan and Horvitz, 1993): accessory neurons, which facilitate this reaction, are missing in Strongyloides. Rhabdias bufonis, a sister taxon to (Strongyloididae 1 Rhabditophanes), displays similar modes of entry and migration. Steinernema primarily enters through body orifices such as malphigian tubules, suggesting that it possess anterio-radial mechanoreception similar to Strongyloides.

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4.3. Does Rhabditophanes represent a reversal of parasitism in Strongyloidoidea? The free-living Rhabditophanes sp. KR3021 was found to be more closely related to Strongyloididae than Rhabdias. As outlined above, Rhabditophanes is free living by all measures: it has continuing fecundity in serial culture, and has never been described from a parasitic association. Two models are possible. Either vertebrate parasitism arose once within the group and was lost in the lineage leading to Rhabditophanes, or parasitism arose twice, in Rhabdias and Strongyloididae. In general, the high degree of specialisation involved in parasitism would suggest that this shift in trophic behaviour is irreversible and that structures and functions once lost are not likely to be regained. However, by establishing a life history with both within-host and inenvironment cycles, Strongyloidoidea have retained the genetic capacity for both free living and parasitic lifecycles. It is possible that within the lineage leading to Rhabditophanes, the capacity for parasitism has been lost. If parasitism arose twice in the group then it must be assumed that many shared parasitic phenotypes also arose independently within Strongyloididae and Rhabdiasidae, which is highly improbable. A single origin for parasitism is more likely, and under this hypothesis Rhabditophanes represents the first known example of reversal of parasitism in a metazoan (Fig. 1). Rhabditophanes is classified within the Alloionematidae (De Ley, 1995). The only other genus within Alloionematidae is Alloionema (not sampled here), a parasite of slugs and snails. Alloionema may have several free-living cycles (Cabaret and Morand, 1990). Free-living cycles may have become extended in the absence of a host reservoir until the free-living life history was fixed. The hypothesis of reversal may be tested by examination of the anterior sensillae of Steinernema, Rhabdias or Rhabditophanes, and SSU rDNA analysis of additional Alloionematidae and the other two described Parastrongyloides spp. The sphaerularoid nematodes, parasites of insects, also display lifecycles, which include alternating generations, reminiscent of the patterns displayed by the Strongyloidoidea. These nematodes (there are 208 species in 29 genera including Allantonema, Heterotylenchulus, Sphaeruliaria, Beddingia and Fergusobia) include all the insect-parasitic tylenchids (Nickle, 1991; Siddiqi, 1981). Tylenchida is included in Clade IV along with the cephalobids and the strongyloidoids. It will be very interesting to examine the species from these insect parasites to more accurately place their radiation with respect to the vertebrate parasites. 4.4. The recent radiation of Strongyloides in vertebrates Parastrongyloides was the closest taxon to Strongyloides spp., as was predicted from morphological considerations. Parastrongyloides has parasitic males and can undergo multiple free-living cycles. This may be the ancestral state

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for the vertebrate parasites given the amphimictic reproductive strategy of Steinernema and other cephalobids. In Strongyloides and Rhabdias, parthenogenesis and hermaphroditism have taken over, respectively, presumably to enable a clonally derived increase in brood size. The genetic similarity among Strongyloides spp. (see Fig. 4a) is striking, given that the branch lengths supporting the genus and close relatives are the longest within any nematode group (Fig. 3 and Blaxter et al., 1998) and that the region chosen for analysis contains the most variable regions of the SSU rDNA gene. This strongly suggests a recent radiation of Strongyloides parasites. Excluding S. westeri, the genus contains two clades. The first of these (the ‘stercoralis’ clade: Strongyloides sp. ex snake ,S. stercoralis and S. f. fuelleborni) can be distinguished from the second (‘cebus’: S. ratti, S. suis, S. venezuelensis, S. papillosus, S. f. kelleyi and S. cebus) by a putative synapomorphic secondary structure region within the E9-2 hairpin of the V2 variable region (Fig. 5). When mapped onto this molecular phylogeny, some aspects of morphology are seen to undergo frequent changes. For example, the arrangement of the ovaries with respect to the gut is an important character for morphological systematics of the group. Within the ‘stercoralis’ clade, S. stercoralis has straight ovaries and Strongyloides sp. ex snake (if identified as S. serpentis) has ovaries that are anteriorly spiral and posteriorly straight, but the ovaries are spiral in S. f. fuelleborni. In the ‘cebus’ clade, S. ratti displays a straight ovary phenotype, but all the other members have spiral ovaries, present in all subsequent lineages. Strongyloides westeri has spiral ovaries. As with other features of nematode morphology, genetic analysis in C. elegans and other related nematodes has shown that such gross variation can be effected by simple (often single) genetic changes (Delattre and Felix, 2001, Felix, 1999, Felix et al., 2000, Fitch and Emmons, 1995, Sommer, 1997). This may also be true for those morphological traits within Strongyloides. Two of the three major causative agents of human strongyloidiasis are grouped together. It is surprising that Strongyloides from a snake forms a sister taxon to the primate parasites. A primate origin for this particular snake house infection can be ruled out due to the distinct E-9 hairpin loop regions of the taxa in this group (see Fig. 5). When derived from another captive snake, the most likely source of the snake Strongyloides would be pit-vipers (Natrix spp.), from which only two-snake Strongyloides spp. have been described as pit-vipers are present in the same snake house from which the Strongyloides analysed herein was sampled. In contrast, while S. ratti and S. venezuelensis are in the same clade, they are significantly separated genetically (mostly through apomorphic changes in the S. venezuelensis SSU sequence) despite inhabiting the same hosts. Strongyloides fuelleborni kelleyi was not supported as a subspecies of S. f. fuelleborni, and thus should perhaps be elevated to specific rank (as S. kelleyi). A likely explanation

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for the origins of S. f. kelleyi strongyloidiasis in New Guinea humans is via a zoonotic source. Strongyloides cebus can be ruled out as there are no non-human primates in New Guinea. The sequences determined from S. papillosus and S. f. kelleyi were identical, suggesting that the human infection arose from S. papillosus, which parasitises a wide range of domesticated animals. The sequence identity is unlikely to be due to contamination, as the amplifications yielding the SSU fragments were performed independently separated by several months. If human strongyloidiasis is derived from S. papillosus then the parasites must have been transferred from domesticated animals to New Guinea humans less than 5,000 years ago, around the time these animals were brought to New Guinea (Viney et al., 1991). The use of S. ratti as an experimental model for human-parasite disease processes would appear to be justified in terms of studying the parasitic phenotype. However, Rhabditophanes may provide an excellent genotypic model given its ease of culture and free-living lifecycle. This study found no support for strong co-evolution between host and parasite. Rather an explosive radiation and host colonisation or switching are indicated. Establishing a time axis for parasite radiation is not possible by direct paleontological comparison but recent radiation is suggested by the genetic similarity within the genus. There is little molecular correlation with morphological features of Strongyloides and thus this proposed phylogeny differs from that predicted from morphological and ecological criteria alone. Acknowledgements We would like to thank colleagues who have supplied us with materials: Drs Tom Nutman, Tom Moore, Gerry Schad, Alex Reid, Einhardt Schierenberg and Warwick Grant. The Rhabditophanes strain was supplied by Dr Kelley Thomas, and The Caenorhabditis Genetics Centre supplied C. elegans. We would also like to thank members of the Blaxter laboratory for support and discussion, and Dr D. Arnot for use of the PCR flow hood. This work was funded through an NERC doctoral fellowship to M.D., and a grant from the Linnaean Society of London to M.B. References Anderson, R.C., 2000. Nematode Parasites of Vertebrates. Their Development and Transmission, C.A.B. International, Wallingford. Ashford, R.W., Barnish, G., 1989. Strongyloides fuelleborni and similar parasites in animals and man. In: Grove, D.I. (Ed.). Strongyloidiasis: A Major Roundworm Infection of Man, Taylor and Francis, London, pp. 271–87. Ashton, F.T., Bhopale, V.M., Fine, A.E., Schad, G.A., 1995. Sensory neuroanatomy of a skin-penetrating nematode parasite: Strongyloides stercoralis. J. Comp. Neurol. 357, 281–95. Ashton, F.T., Li, J., Schad, G.A., 1999. Chemo- and thermosensory neurons: structure and function in animal parasitic nematodes. Vet. Parasitol. 84, 297–316.

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