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Three Hoplolaimus stephanus populations were characterized morphologically, both by morphometrics and by SEM observations. These populations were used to develop a rapid and accurate molecular identification method for the species, which is useful because of the high level of morphological similarity between H. stephanus and H. galeatus. Species-specific primers for H. stephanus, amplifying a distinct fragment (260 bp) of the nuclear ribosomal internal transcribed spacer1 (ITS1), can be used in multiplex PCR along with previously developed primers for other common Hoplolaimus species. We also infer phylogenetic relationships among H. stephanus, the closely-related H. galeatus, and several other Hoplolaimus species, using sequences of the actin gene, ITS1 and LSUD.
Hoplolaimus stephanus Sher, 1963 was first described from swamp soil in Nicols, South Carolina, USA (Sher, 1963). Vovlas et al. (1991) later provided additional scanning electron microscopy (SEM) observations and supplementary morphometric data from a population collected in Raleigh, North Carolina, USA. The host plants were not identified in either of these two publications, which are also the only two published references available for this species.
We identified three populations of H. stephanus, one in Pennsylvania and two in South Carolina, USA. To ensure accurate identification, combined morphological and molecular analyses were initiated. H. stephanus morphologically resembles H. galeatus, but it can be distinguished from H. galeatus by the 24 to 28 longitudinal striations on the basal annule of the lip region compared to 32-36 in H. galeatus, shorter spicules, less areolation of the lateral field, and shorter body length (Sher, 1963). There are no publicly available DNA sequences for H. stephanus and this species has not been genetically characterized.
Hoplolaimus spp. reported from the southeastern USA include H. columbus Sher, 1963, H. galeatus (Cobb, 1913) Thorne, 1935, H. magnistylus Robbins, 1982, H. stephanus, H. seinhorsti Luc, 1958 and H. tylenchiformis von Daday, 1905(Lewis and Fassuliotis, 1982). H. columbus, H. galeatus and H. magnistylus, are considered to be economically important and can cause serious damage to agronomic crops, including cotton (Gossypium hirsutum L.), corn (Zea mays L.) and soybean (Glycine max L.) (Fassuliotis, 1974; Nyczepir and Lewis, 1979; Robbins et al., 1987, 1989; Henn and Dunn, 1989; Noe, 1993).
Hoplolaimus galeatus feeds in both the cortex and vascular tissues in cotton and causes extensive damage in the vascular cylinder (Krusberg and Sasser, 1956). It is also a prevalent pathogen of turf grasses such as St. Augustinegrass (Stenotaphrum secundatum) and bermudagrass (Cynodon dactylon) in Florida (Henn and Dunn, 1989; Giblin-Davis et al., 1990, 1995). It has a wide distribution range in the United States (Wrather et al., 1992; Lewis et al., 1993; Martin et al., 1994; Gazaway and McLean, 2003), and there are records from Canada, South America, Central America, and India on a variety of hosts (Fortuner, 1991). Because the morphometric values of H. stephanus and H. galeatus overlap considerably (Sher 1963; Vovlas et al., 1991; Handoo and Golden, 1992) and because the useful morphological characters for their discrimination require high magnification and considerable diagnostic time, it is important to improve our ability to accurately diagnose these species. Our finding H. stephanus populations associated with two woody hosts, birch tree and dogwood tree (Betula nigra L. and Cornus florida L.) and one grass host (Poa pratensis L.) indicates that the host range for H. stephanus overlaps that of H. galeatus, and evidences overlap in the geographic distribution as well.
Species identification is a prerequisite to developing resistant varieties and to determining basic biology and behavior of lance nematodes. Molecular diagnostic techniques based on sequence variation in the ITS region have been developed to discriminate the species in Hoplolaimus. Species-specific primers to identify H. columbus, H. galeatus and H. magnistylus were recently developed by Bae et al. (2009). A number of ITS and LSU sequences are publicly available in GenBank for these and a few more species of Hoplolaimus, but not for H. stephanus. Our objectives were to develop a rapid and accurate diagnostic molecular identification method for H. stephanus and to infer phylogenetic relationships among this species and the closely-related H. galeatus.
Nematode samples and DNA extraction: The species used in this study are listed in Table 1, along with the geographic origin and host plant of the populations. Three populations of H. stephanus, including one from Dillsburg, Pennsylvania (on Kentucky bluegrass) and two from the campus of Clemson University, Clemson, South Carolina (one on dogwood and one on river birch) were used for the morphological and molecular characterizations. Two populations of H. columbus from South Carolina (one from a cotton field and one from a soybean field), and two populations of H. galeatus from South Carolina (one on St. Augustine grass and one on bermudagrass) were also included in the analyses. Forty-seven original sequences generated in this study were added to GenBank, but publicly available sequences from other authors for several Hoplolaimus species and two outgroup species were included for comparison. The accession numbers for all sequences included in this study are listed in Table 1.
DNA was extracted from individual nematodes hand-picked from each population, using Sigma Extract-N-Amp kit (XNAT2) (Sigma, St. Louis, MO). The manufacturer's protocol was modified by reducing all volumes to one eighth of the recommended amounts. One nematode was placed into a 0.2 ml centrifuge tube containing 12.5 μl of the kit's Extraction Solution. The nematode was then crushed using the tip of a <10 μl pipette tip, followed by adding 3.5 μl of the kit's Tissue Prep solution to the tube. The tube was then vortexed, followed by a brief centrifugation to collect contents. Tubes were then incubated at 55 °C for 10 minutes, followed by incubation at 95 °C for 3 minutes. Next, 12.5 μl of the kit's Neutralization Solution was added to each tube. The extracted DNA was used for PCR or stored at -20°C.
Morphological characterizations: At least 8 females and 4 males from each population were used for the morphological diagnosis. Measurements and observation of morphological diagnostic characters were made using temporary mounts in water and with an Olympus BX60 microscope equipped with the software iSolution Lite (Image and Microscope Technology i-Solution, Inc.).
Scanning electron microscopy (SEM) observations were conducted on a Hitachi Analytical Tabletop Microscope TM3000. Male and female specimens of H. stephanus and H. galeatus were fixed in 2.5% glutaraldehyde for at least 2 hours, then passed through a graded series of ethanol dehydration (25%, 50%, 75%, 90%, 95% , 100% ethanol, 15 min each), followed by critical point drying with hexamethyldisilazane (HMDS) and platinum coating prior to examination.
Molecular characterization: Molecular characterization of populations was conducted using species-specific primers and methodology as described by Bae et al. (2009) for discrimination of H. galeatus, H. columbus, and H. magnistylus.
H. stephanus species-specific primer design: The species-specific primers for H. stephanus were designed using comparative ITS1 region sequence alignments (Fig. 1) of Hoplolaimus columbus, H. galeatus, H. magnistylus, H. seinhorsti, H. concaudajuvencus and H. stephanus (gi226431011, gi186920200, gi186920202, gi186920203, gi186920204, H. stephanusSCST01). Five putative sets of primers were designed with the Primer-Blast application on NCBI (http://www.ncbi.nlm.nih.gov/tools/primer-blast). After testing under a range of annealing temperatures (54-61°C), species-specific primer pair Hs-1f (5’- CCTGCCTTGGGGGTCGCTTG-3’) and Hs-1r (5’- GCCAGTGTGTTCCGCTCGCA-3’) were chosen and optimal PCR reactions with these primers were performed with annealing temperature of 60°C.
PCR Amplification of actin, ITS1 and LSUD genes: The actin region was amplified by the primers Act1-f (5’-CCAAATCATGTTCGAGACGTT-3’) and Act1-r (5’-GAACATAGCCTCTGGGCAAC-3’). These primers were designed using comparative sequence alignments of Heterodera cynodontis, Heterodera avenae, Heterodera schachtii, Heterodera latipons, Heterodera glycines and Caenorhabditis elegans from GenBank (Accession numbers gi167472950, gi41387727, gi41387723, gi41387721, gi26422171, gi18314322, gi133952034). The best putative primers were selected using the Primer-Blast application of NCBI (http://www.ncbi.nlm.nih.gov/tools/primer-blast). Amplifications were performed in 20μl reactions, each containing: 5 μl PCR-grade water, 10 μl of ReadyMix TAq PCR Reaction Mix with MgCl2 (Sigma, St. Louis, MO) (20 mM Tris-HCl pH 8.3, 100 mM KCl, 3 mM MgCl2, 0.002% gelatin, 0.4 mM dNTP mixture (dATP, dCTP, dGTP, and dTTP), and 0.06 units of Taq DNA Polymerase/μl), 1.0 μl of each primer (20 μM), and 2 μl of DNA template. The PCR reactions were performed on a PTC-100 Peltier thermal cycler (MJ Research, Inc. Watertown, Massachusetts) with the following run parameters: one initial denaturation cycle at 95°C for 3 min, followed by 36 cycles at 95°C for 45s, 46°C for 1.5 min, 72°C for 2 min and final extension at 72°C for 10 min.
The ITS1 and LSU-D region were amplified using the primers Hoc-1f (5’-AACCTGCTGCTGGATCATTA-3’), Hoc-2r (5’-CCGAGTGATCCACCGATAA-3’), LSUD-f (5’-ACCCGCTGAACTTAAGCATAT-3’) and LSUD-r (5’-TTTCGCCCCTATACCCAAGTC-3’), respectively, designed by Bae et al. (2008). PCR reactions were performed as described above, using the following parameters: an initial denaturation cycle at 95°C for 3 min, 36 cycles at 95°C for 45s, 59°C for 1.5 min, 72°C for 2 min, and a final extension cycle at 72°C for 10 min. Each reaction included a negative control without DNA template. After amplification, 5 μl of each reaction were loaded onto a 1.5% agarose gel (120 V, 50 min) and photographed under UV light. At least three replicates were performed for each population-primer set combination. PCR products were purified and concentrated with Bio-Rad PCR Kleen Spin Columns (Bio-Rad, Hercules, CA). Purified DNA was sent to the Clemson University Genomics Institute (Clemson, SC) for direct sequencing in both directions. Amplification primers were used as sequencing primers. The sequences were edited and aligned using BioEdit 7.0 (Hall, 1999).
Phylogenetic analysis: Sequences of ingroup and outgroup taxa were aligned using ClustalW (Thompson et al., 1994). Sequences of Globodera rostochiensis and Heterodera glycines were used as the outgroup taxon for actin, ITS1 and 28S region (Table 1). The files were converted from FASTA to NEXUS format using DNA Sequence Polymorphism Version 5.10.01 (DnaSPv5) (Librado and Rozas, 2009). A best fit model of nucleotide substitution was selected using the GTR+I+G model with the Akaike Information Criterion (AIC) among 56 different models using ModelTest v 3.7 (Posada and Crandall, 1998). The Akaike-supported model, the base frequencies, the proportion of invariable sites, and the gamma distribution shape parameters and substitution rates were used in phylogenetic analyses. Bayesian inference was implemented for each gene separately using MrBayes 3.1.2 program (Huelsenbeck and Ronquist, 2001) running the chain for 100,000 generations with the Markov Chain Monte Carlo (MCMC) method, a sample frequency of 10 and burn-in value of 250. We estimated the posterior probabilities of the phylogenetic trees (Larget and Simon, 1999) using the 50% majority rule. The phylogenentic trees were viewed with TreeView1.6.6 (Page, 1996).
Morphological characterization: The measurements and diagnostic morphological characters for the three H. stephanus populations included in this study are presented in Table 2. We include a comparison to the measurements and characters of H. stephanus published by Sher (1963) and by Vovlas et al. (1991), and a comparison to morphometrics and characters of a H. galeatus population published in Sher's (1963) and Handoo and Golden's (1992) revisions of the genus. Our measurements and morphological characters for the three populations conform to the original description of H. stephanus, except for the larger size of females (1.28-1.67mm vs 1.01-1.45mm) of the dogwood population. The grass and birch populations were very similar to Sher's (1963). We found greater differences with the observations by Vovlas et al. (1991), who report more longitudinal striations on the basal lip annule (30-36), and also larger variation of the number of tail annules (11-15). Our observations suggest that tail annules (12) are a stable character, as are the number of lip annules (4).
Our observations confirm the diagnostic characters for H. stephanus, including four lip annules (Fig. 2A), basal lip annule with 24-28 longitudinal striae (Fig. 2A), presence of males (Fig. 2B), four lateral incisures (Figs. 2C-F), presence of epiptygma (Fig. 2E), and incomplete areolation of the lateral field (Fig. 2E, 2F). Our observations on the presence of epiptygma coincide with those reported by Volvas et al. (1991).
Molecular identification: Multiplex PCR products amplified from H. columbus, H. galeatus and H. stephanus templates using our H. stephanus species-specific primer set and resolved using agarose gel electrophoresis are shown in Figure 3. An internal positive control for each reaction was included by using primers LSUD-1f and LSUD-2r (amplicon size of 1030bp). Primers HS-1f and HS-1r do not amplify H. columbus (lanes 1-3) or H. galeatus (lanes 4-6). These primers only amplify H. stephanus (lanes 7-9, 260bp fragment).
The species-specific primers designed by Bae et al. (2009) accurately discriminate H. columbus, H. galeatus and H. magnistylus. In Figure 4, multiplex PCR products amplified from H. columbus, H. galeatus and H. stephanus templates using these species-specific primer sets are shown. An LSUD fragment was amplified as internal positive control for each reaction. Lanes 1-3 show H. columbus (580 bp fragment), lanes 4-6 show H. galeatus (120 bp fragment), and lanes 7-9 show H. stephanus (amplification of only the internal control).
Phylogenetic analysis: All three H. stephanus populations considered in this study were grouped together in Clade I (Figures 5-7), with high posterior probability (100% in ITS1 and actin trees, 99% in LSUD tree), which supports our morphological identification and characterization of the populations. The unknown species Hoplolaimus sp.3, from GenBank and Bae et al. (2008) was grouped into the same clade with H. stephanus in both ITS1 and LSUD phylogram trees. The BLAST search of the sequences of both ITS1 and LSUD fragments from the three populations of H. stephanus in this study result in high sequence similarity (99%) with this unknown species. We are confident that the specimens reported as unknown species 3 are H. stephanus.
Unknown species Hoplolaimus sp.1 and Hoplolaimus sp.2 from Bae et al. (2008) are very close to H. stephanus (96% and 98% sequence similarity, respectively), but likely different species. In silico analysis indicate that our proposed H. stephanus-specific primers will not amplify these two species, but actual specimens will be required to validate this and to characterize the species morphologically. Until we are able to obtain more of these specimens, these two species will remain uncertain.
Phylogenetic analysis of Bayesian Inference using actin, ITS1 and LSUD sequences produced phylogram trees with similar topological relations among the Hoplolaimus species. The actin gene tree in Fig.5 indicates a close relationship between H. stephanus and H. galeatus. The LSUD tree in Fig. 6 groups H. columbus and H. seinhorsti into the same Clade II; and H. magnistylus, H. galeatus, H. concaudajuvencus and H. stephanus into a big Clade III, supported by 0.81 posterior probability. These two clades coincide with the proposal by Siddiqi (2000) of divisions into two subgenera: i) subgenus Hoplolaimus Daday, 1905 that includes species with lateral field with four incisures, such as H. galeatus, H. stephanus, H. concaudajuvencus, and H. magnistylus; and ii) subgenus Basirolaimus Shamsi, 1979 that includes species with one to three incisures, such as H. columbus and H. seinhorsti.
H. stephanus is more commonly occurring than currently reflected in the literature, and this leads us to believe that it is very probable that some reports of H. galeatus could in fact be H. stephanus, especially considering the overlap in host plants and geographic distribution. Our phylogenetic analysis supports H. stephanus as a valid phylogenetic species that can be easily discriminated genetically. However, it can be challenging to differentiate H. galeatus from H. stephanus by morphology. Our species-specific primers will therefore prove very useful for identification.
The actin gene has been widely used in phylogenetic analysis of other nematode groups, like Globodera and Heterodera species (Kovaleva et al., 2005). We believe that using this gene together with the ribosomal sequences will increase the robustness of phylogenetic analyses of relationships among Hoplolaimus species. In this study, the application of three genetic markers provided a higher resolution analysis of taxonomic relationships and better support for the delimitation of H. stephanus (Figure 5-7).
It is necessary to undertake studies that will compare the ecology and biology of H. stephanus and H. galeatus. Pathogenicity studies on potential common hosts, such as grasses and woody species, could reveal the need to develop different threshold levels for these two species. Edaphic factors studies, mainly those concerning observations on correlations with soil texture and levels of disturbance, could help predict the presence and geographic distribution of these two closely-related species.
This research was supported by South Carolina Soybean Board.
This paper was edited by Dan Bumbarger.