Figures
Abstract
The beet cyst nematode, Heterodera schachtii, is a major root pest that significantly impacts the yield of sugar beet, brassicas and related species. There has been limited molecular characterisation of this important plant pathogen: to identify target genes for its control the transcriptome of the pre-parasitic J2 stage of H. schachtii was sequenced using Roche GS FLX. Ninety seven percent of reads (i.e., 387,668) with an average PHRED score > 22 were assembled with CAP3 and CLC Genomics Workbench into 37,345 and 47,263 contigs, respectively. The transcripts were annotated by comparing with gene and genomic sequences of other nematodes and annotated proteins on public databases. The annotated transcripts were much more similar to sequences of Heterodera glycines than to those of Globodera pallida and root knot nematodes (Meloidogyne spp.). Analysis of these transcripts showed that a subset of 2,918 transcripts was common to free-living and plant parasitic nematodes suggesting that this subset is involved in general nematode metabolism and development. A set of 148 contigs and 183 singletons encoding putative homologues of effectors previously characterised for plant parasitic nematodes were also identified: these are known to be important for parasitism of host plants during migration through tissues or feeding from cells or are thought to be involved in evasion or modulation of host defences. In addition, the presence of sequences from a nematode virus is suggested. The sequencing and annotation of this transcriptome significantly adds to the genetic data available for H. schachtii, and identifies genes primed to undertake required roles in the critical pre-parasitic and early post-parasitic J2 stages. These data provide new information for identifying potential gene targets for future protection of susceptible crops against H. schachtii.
Citation: Fosu-Nyarko J, Nicol P, Naz F, Gill R, Jones MGK (2016) Analysis of the Transcriptome of the Infective Stage of the Beet Cyst Nematode, H. schachtii. PLoS ONE 11(1): e0147511. https://doi.org/10.1371/journal.pone.0147511
Editor: Philippe Castagnone-Sereno, INRA, FRANCE
Received: December 30, 2014; Accepted: January 5, 2016; Published: January 29, 2016
Copyright: © 2016 Fosu-Nyarko et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All reads used for the transcriptome analysis have been deposited at NCBI Sequence Read Archive under the accession SRX381021.
Funding: Murdoch University provided PhD studentship for PN; RG was supported by an Australian India Strategic Research Fund Grant (BF030027); Nemgenix Pty. Ltd. contributed funds for consumables and support in the form of salaries for JFN, but did not have any additional role in the study design, data collection and analysis, decision to publish, or preparation of the manuscript. The specific roles of this author is articulated in the "author contributions" section.
Competing interests: Author JFN was employed by Nemgenix Pty. Ltd at the time of review of the manuscript. Nemgenix Pty. Ltd contributed funds for consumables. There were no patents, products in development or marketed products to declare. This does not alter the authors' adherence to all the PLOS ONE policies on sharing data and materials.
Introduction
The beet cyst nematode, H. schachtii, is a sedentary endoparasitic plant nematode present in temperate and mediterranean regions. It has a narrow host range but can substantially impact infected crops, with losses of 30% common in hosts of the families Chenopodiaceae (especially Beta vulgaris) and Cruciferae [1, 2]. Heavy root infestation with H. schachtii can cause root distortion, ‘bearding’ of sugar beet storage roots and lateral root death: plants become prone to wilting and there is a reduction in plant growth and yield. Plant parasitic nematodes (PPNs) are usually managed by a combination of crop rotation, application of nematicidal agrochemicals and deployment of natural resistance genes, and each of these has limitations. In particular, nematicides are either too expensive, toxic over the longer term, some have been banned or their use restricted, and there are few non-hosts to use as break crops in crop rotations. In many cases resistant genes are not available or effective against PPNs [3–5]. These factors make the search for alternative control methods for nematodes more imperative. H. schachtii is of additional interest because it is one of the few PPNs which can infect Arabidopsis thaliana, the best understood model plant species [6].
Pre-parasitic cyst nematode stage 2 juveniles (J2s) moult from J1s that develop from eggs within the cysts, and then migrate to host plants (Fig 1). They then enter root cells using their hollow mouth stylet, aided by secretions produced from gland cells that include cell wall-modifying enzymes such as cellulases and pectinases, and migrate intracellularly, responding to positional gradients in the root [7–9]. In a susceptible host, the nematode becomes established after forming a typical feeding site (syncytium). The syncytium forms through dissolution of cell walls and cell expansion to create a multinucleate region of interconnected and metabolically active cells, from which it feeds throughout its life cycle [10–12]. Secretions from J2s are required for root entry and migration, and in initiation and establishment of a syncytium by co-ordinated modification of expression of a series of host genes. Once a syncytium has been initiated successfully, J2s become sedentary, and each feeds from its associated syncytium, growing and developing via three further moults to maturity. Males are associated with smaller syncytia—they emerge from roots, and fertilise the females. Adult females become lemon-shaped and after fertilisation, produce many eggs, most of which are retained within the body. When the female nematode dies the body wall tans and hardens to form a protective cyst around the eggs.
Genetic studies of H. schachtii have been limited so far to targeted isolation of transcripts/genes of putative effectors to study the parasitome (i.e. secreted products of parasitism genes) by direct amplification, cloning and sequencing [8, 13, 14]. Recent advances in sequencing technology and bioinformatic analysis platforms have led to increased understanding of nematode biology. For example, the genomic sequencing and characterisation of genes of the free-living Caenorhabditis spp. has contributed to annotation of sequenced genomes of animal parasitic nematodes and PPNs such as the root knot nematodes Meloidogyne hapla and Meloidogyne incognita [15, 16], the cyst nematode G. pallida [17] and recently the migratory endoparasitic nematode Pratylenchus coffeae [18]. In addition, global transcriptomic methods allow complex sets of genes and pathways in an organism to be analysed simultaneously, so that in the case of nematodes it can be used to study the expression of genes at different life stages, including genes involved in parasitism (e.g. [19–22]). At present there are only 3,066 nucleotide sequences for H. schachtii in the largest sequence database, the National Centre for Biotechnology Information (NCBI) Genbank (http://www.ncbi.nlm.nih.gov/) and 1,592 on nematode.net (http://nematode.net), and a global transcriptome analysis has not been published so far for any stage of the nematode.
In this paper, we report the sequencing of the transcriptome of the infective J2 stage of H. schachtii prior to root entry, and annotation of transcripts using sequences of both free-living and parasitic nematodes, and conserved or core eukaryotic genes. Several bioinformatic tools were used to identify a set of genes that may be primed for host infection. The infective J2 stage appears to express many of the genes required for migration through host cells and for initiation of syncytia. Transcripts with homologies to the recently identified soybean cyst nematode nyavirus were also found [23]. Analysis of the transcriptome of J2s of H. schachtii provides an additional resource for functional genetic analysis of this PPN. Although many of the identified genes need further functional characterisation, it also provides a set of sequences that could be used to exploit gene-based strategies such as RNA interference, to develop new strategies for control of this economically important nematode.
Materials and Methods
Ethics Statement
The population of H. schachtii used for the study was originally obtained from a cabbage and broccoli farm north of Perth, Western Australia. Permission to enter the farm was granted by the farm owner who also kindly provided soil samples from a freshly harvested plot of land from which we obtained mature female cysts. No specific regulatory permissions were required for these locations/activities because H. schachtii is an endemic pest in Western Australia, and these nematodes are not protected or an endangered species.
Transcriptome sequencing and assembly of reads
The population of H. schachtii used for transcriptome sequencing was derived from a single female and was maintained on cabbage plants grown in white sand in a glasshouse at 25°C (day) and 15°C (night). Pre-parasitic J2s were harvested using a mist apparatus described by Tan et al [24] (S1 Fig). Using this method, it was expected that the pre-parasitic J2s extracted, which had been exposed to roots of susceptible hosts, were primed for infection, and the genes expressed at this stage were analysed. For J2 nematode isolation, soil taken from the root zone of infected plants was placed in a 200 mL plastic container lined with two layers of coffee filters. The soil was sprayed with a water mist every 10 minutes for 10 seconds, and active nematodes migrated through the double coffee filter and were collected at four hourly intervals in aerated fresh water that was continually gently replaced to remove microbial contamination (S1 Fig). To further prevent contamination, the active freshly extracted nematodes were then surface-sterilised by suspension in 1% chlorhexidine gluconate (hibitane) for 20 min, followed by washing with 1% streptomycin for 2 min and then three washes with sterile distilled water: for each treatment and wash the nematodes were harvested by gentle centrifugation at 1200 g for 2 minutes. The J2s were then examined by light microscopy to ensure they were viable and free from any visible microbial contamination before RNA extraction. Total RNA was extracted with Trizol (Invitrogen Life Technologies, Carlsbad, USA) and cleaned using a RNeasy Mini kit column (Qiagen Pty. Ltd., Victoria, Australia). The quality and quantity of RNA was assessed with an Agilent 2100 Bioanalyzer (Agilent Technologies, Mississauga, Canada) and RNA with a minimum RNA Integrity Number (RIN) of 7 and with absorbance ratios of 260:280 at 2.0 and 260:230 at 2.1 were used for cDNA synthesis. A cDNA library was prepared from 3 μg of total RNA using the Ovation RNA-Seq system (NuGEN Technologies Inc., CA, USA), which uses both Oligo-dT and random primers, and checked for removal of rRNA using an Agilent 2100 Bioanalyzer. Sequencing (half a picotitre plate) was carried out using a Roche 454 GS FLX DNA platform at the Institute of Immunology and Infectious Diseases, Murdoch University, Perth, Australia.
Reads that passed the key filter test based on default parameters for base calling were trimmed and analysed after quality control. The average PHRED scores for the reads (mean of PHRED scores for bases making up an entire read) were then determined using the CLC Genomics Workbench 7.0.4 after which high quality reads were assembled with the CAP3 assembler and the CLC Genomics Workbench 7.0.4. CAP3 was configured to run with an overlap per cent identity cut-off of 90, overlap length cut-off of 40, mismatch score factor of -5 and other default settings [25]. The set of parameters used for the CLC Genomics Workbench assembly were: mismatch cost of 2, insertion cost of 2, deletion cost of 2, length fraction of 0.4 and similarity fraction of 0.4.
Annotation and functional classification of transcripts
For most analyses TBLASTX 2.2.30+ [26] was used to compare transcripts to reference sequences in defined NCBI databases or local databases (e.g. created for CLC Genomics Workbench 7.0.4). All BLAST analyses were done with a threshold e-value of 1E-05 after which hits with significant identities were sorted or manually curated using High-scoring Segment Pair (HSP) scores or total bit scores as these parameters are independent of the search space size and enable comparisons across databases and searches. Multiple sequence alignments of transcripts and matches presented were done using BioEdit [27]. Transcripts were annotated based on similarities with sequences in the NCBI non-redundant (nr/nt), Gene Ontology (GO, submission date (30/9/2014, geneontology.org/page/download-annotations) [28], and Carbohydrate Active Enzyme (CAZy, November, 2014 release, www.ahv.dk) [29] databases. In particular, they were compared to protein sequences of well-characterised free-living nematodes (Caenorhabditis spp.) to determine genes expressed in the H. schachtii J2 transcriptome and their putative functions using the GO classification, based on C. elegans genes. Transcripts for proteins involved in RNA interference (RNAi) pathways were also identified using those characterised for C. elegans: these proteins were identified from the literature and from descriptions on Wormbase (www.wormbase.org): their amino acid sequences were downloaded and used as a reference for BLASTX searches. Term identifiers for PAMGO, Plant Associated Microbe Gene Ontology, were used to determine association of transcripts with known plant host-pathogen interactions [30].
Comparative analyses with ESTs and genes of other nematodes
The H. schachtii J2 transcriptome was compared to genes and ESTs of a reference group of 17 nematode species available at NCBI using a local TBLASTX/BLASTX with a threshold e-value of 1E-05. The reference nematode groups used were the free-living nematodes C. elegans and Caenorhabditis remanei, and three PPN groups with different modes of feeding: cyst nematodes (H. glycines, H. schachtii, G. pallida, Globodera mexicana, Globodera rostochiensis), the root knot nematodes Meloidogyne spp: M. arenaria, M. chitwoodi, M. javanica, M. paranaensis, M. incognita, M. hapla) and migratory nematodes (Pratylenchus vulnus, Pratylenchus penetrans, Pratylenchus thornei and Radopholus similis). The numbers of ESTs, genomic contigs and protein sequences for these nematode species as well as sequences in databases used for functional annotation (GO, CAZy, PAMGO) are provided in S1 Table.
Mapping H. schachtii transcriptome to genomes of four PPNs
The H. schachtii transcriptome was also compared with publicly available genomic contigs of two cyst [H. glycines (PRJNA28939) and G. pallida (PRJEB123)] and two root knot nematodes [M. hapla (PRJNA29083) and M. incognita (PRJEA28837)], both directly and indirectly in four ways. BLASTN and TBLASTX were first used separately to study relatedness of the transcripts to those for putative proteins encoded by the genomic sequences. The H. schachtii reads were also mapped onto the genomic contigs of each of the four nematodes using the “map to reference” function of CLC Genomics Workbench 7.0.4 with a mismatch cost of 2, an insertion cost of 3, a deletion cost of 3, a length fraction of 0.5 and a similarity fraction of 0.8. Finally, genes expressed at the pre-parasitic stage were assessed using the Core Eukaryotic Genes (CEGs) used in CEGMA (Core Eukaryotic Genes Mapping Approach [31]), and were also compared to CEGs encoded by genomic sequences of the four plant parasitic nematodes.
Identification of spliceosome genes and evidence of trans-splicing in H. schachtii
Further analysis was conducted with the transcriptome to study genes involved in the spliceosome of cyst nematodes, in particular, the phenomenon of trans-splicing. H. schachtii orthologues of such genes were identified using the KEGG spliceosome pathway of C. elegans as a reference (www.genome.jp/kegg). Evidence of trans-splicing in H. schachtii was studied using the presence of splice leader (SL) sequences on transcripts. A SL1 RNA gene was first identified from genomic sequences of H. glycines (PRJNA28939) using BLASTN and sequences for those of C. elegans and Pristionchus pacificus [32] to confirm the presence of such sequences in genomes of cyst nematodes. The identified SL sequence was then compared to trans-splice leaders (SL1-like and SL2-like) of 11 nematode species (including Pristionchus pacificus, Brugia malayi, Ascaris suum, C. elegans and C. briggsae) [33] which identified it as a SL1-like sequence. This conserved SL1-like sequence was then used to identify similar sequences in H. schachtii transcripts.
Putative nematode parasitism effectors of H. schachtii
Transcripts for putative parasitism effectors of H. schachtii were identified using functionally characterised and putative homologues of other PPNs. A local database was created for representative characterised parasitism effectors of cyst, root knot and migratory PPNs and used as a reference for TBLASTX search among the H. schachtii transcripts. The sequences included those for 30 characterised effectors (29 complete coding (mRNA) and one partial cDNA) and all publicly available ESTs isolated from gland cells and secretions of H. glycines and M. incognita. Alignments of best matching transcripts, with the highest bit score, to respective reference sequence(s) were manually curated, for example, to allow identification of the presence of important features such as signal peptides. Also, sequences of transcripts with matches to nematode parasitism effectors and CAZymes were translated into six frames using the translate programme in CLC Genomics Workbench 7.0.4 and the putative proteins of each frame used as a query to search for signal peptides and trans-membrane helices similar to those of eukaryotes using SignalP 4.1 [34].
Results
The transcriptome
The sequencing generated a total of 1,064,983 reads of which 400,622 reads passed key filter tests and were further analysed. From this, 387,668 (97%) high quality reads with a least average PHRED score of 22 were assembled into contigs. From 82% of the high quality reads, the CLC Genomics Workbench generated 47,263 contigs, about 10,000 more than CAP3, which assembled 62% into contigs (Fig 2). Consequently, there were more singletons after the CAP3 assembly, with an average length of 192 nt, which was about 40 bases shorter than those from the CLC Genomics Workbench assembly (Fig 2). The average length and N50 of the CAP3 contigs were both slightly greater: 414 nucleotides (nt) and 477 nt respectively. The total length of the CAP3 and CLC contigs were 35.4% and 19.7%, respectively, of the entire length of all reads of the transcriptome although more reads were assembled by the CLC genomics workbench. For both assemblies, the majority of the contigs were between 300–600 nt long: 60.7% for CAP3 and 52.5% for the CLC Genomics Workbench contigs (Fig 2). The longest contigs for both assemblies were over 4 Kb long (one for each). Generally for the CAP3 assembly there were substantially fewer contigs 100 bases or shorter than from the CLC assembly, with the opposite the case for singletons (Fig 2). Because the CLC Genomics Workbench assembled more reads into contigs and the average length of singletons was longer, the CLC assembly was used for all annotation of the transcriptome. Analysis of available ESTs of other cyst nematodes in the NCBI databases indicate the average length and the N50 for the CLC assembly were both shorter: respectively the average length and N50 for ESTs of the reference nematodes were 368 and 578 for H. schachtii; 267 and 611 for Globodera spp and, 355 and 691 for H. glycines. The generated reads have been deposited in the Sequence Read Archive of NCBI under the accession SRX381021.
A). Size distribution of contigs assembled with CAP3 and CLC Genomics Workbench 7.0.4. B). Size distribution of singletons after read assembly with CAP3 and CLC Genomics Workbench 7.0.4. C). Transcriptome and assembly statistics of CAP3 and CLC Genomics Workbench 7.0.4.
Gene ontology of transcripts
From comparisons with the NCBI non-redundant protein dataset, 87.8% of the annotated transcripts matched eukaryotes whereas 8.9% matched bacterial proteins, 0.5% to viral sequences and the remainder matched archaea, and unclassified sequences. The transcripts matched sequences from 343 species; most of these had high percentage sequence identities to transcripts/genes of Caenorhabditis spp, A. suum, Loa loa and B. malayi. Transcripts from PPNs were relatively under-represented in the results either because there were fewer sequences in databases or the ESTs were shorter than those for the free-living and animal species.
The GO classification scheme based on proteins of Caenorhabditis spp. was used to annotate the transcripts. A total of 6,430 transcripts (3,579 contigs and 2,851 singletons) had similarities to those of 2,530 C. elegans proteins. Using the GO of 2,410 C. elegans proteins for detailed functional classification of the transcripts, 1,915 were involved in biological processes, 42 were associated with cellular components and 1,587 had various molecular functions (Fig 3). A total of 46 H. schachtii transcripts with sequence similarities to those of nine proteins associated with PAMGO terms were identified (S2 Table). These included three proteins for Phytophthora spp and six for Magnaporthe grisea, seven of which were associated with multiple PAMGO terms including the following: ‘pathogenesis’ (GO:0009405), ‘entry into host’ (GO:0044409), ‘chemotaxis’ (GO:0006935), and ‘secretion/toxin transport activity’ (GO:0019534). While some of these terms could be related to activities of a nematode infection process, functions of the associated transcripts and for those associated with molecular terms/functions involved in development (e.g. the GO:0004674-Serine/threonine protein kinase activity) and interaction with a compatible host (e.g. GO:0007165-signal transduction), are yet to be determined for most nematodes.
Comparative analysis with ESTs and genes of nematode species
When the transcripts were compared to genes and ESTs of free-living, sedentary cyst and root knot, and migratory endoparasitic nematodes, those identified with sequence similarity to any of the reference group were collated and presented in a Venn diagram (Fig 4). A total of 33,313 had sequence similarities to ESTs/genes of the reference nematode groups of which 2,918 were common to all the four reference groups of nematodes (Fig 4). Consistent with the number of publicly available reference sequences used for the comparisons, more H. schachtii transcripts matched ESTs of the cyst nematode group (26,464), followed by the root knot group (14,769) and then the free-living (10,518) and the migratory endoparasitic nematodes with 7,083 matched transcripts. The numbers of H. schachtii transcripts with matches to only sequences of each group followed a similar pattern: more for the cyst nematode group, then the root knot, followed by the free living and migratory nematode groups in that order (Fig 4). Most of the 2,918 transcripts which match sequences of all the four group of nematodes putatively encode proteins involved in developmental and common molecular and biological processes of eukaryotes. Not surprisingly, transcripts with no match to any sequence of free-living nematodes included those putatively encoding parasitism effectors.
The transcriptome of H. schachtii and genomes of four PPNs
BLASTN and TBLASTX comparisons revealed 58% of the H. schachtii transcripts mapped to 52% of genomic contigs of the closely related H. glycines at an e-value threshold of 1E-05, whereas in all 43% of the transcripts from both the BLASTN and TBLASTX searches matched 41% of the G. pallida genomic contigs (Fig 5A and 5B). About 20% of the transcripts matched contigs of the root knot nematodes. However, the matching contigs represented close to 63% and 52% of the total genomic contigs for M. hapla and M. incognita respectively (Fig 5A and 5B). Using the ‘map reads to reference’ function on the CLC Genomics Workbench, more (ca 67%) H. schachtii reads were mapped to H. glycines contigs (63%) than to contigs of G. pallida, M. hapla and M. incognita (Fig 5A) and like the BLAST results, the percentage of genomic contigs mapped to the H. schachtii reads were higher for H. glycines, M. hapla, M. incognita than for G. pallida (Fig 5B).
A). Percentage of H. schachtii transcripts with matches to genomic contigs of H. glycines, G. pallida, M. incognita and M. hapla. B). Percentage of genomic contigs of H. glycines, G. pallida, M. incognita and M. hapla with similarity to H. schachtii transcripts. C). Comparison of homologues of CEGs in the H. schachtii transcriptomes to those in genomes of H. glycines, G. pallida, M. incognita and M. hapla.
The 259,111 reads that mapped to the H. glycines contigs had an average length of 192 nt and made up 63.25% of the total bases of the transcriptome. Generally, the number of reads mapping to the genomic contigs did not depend on the read lengths or the composition of the reads since the distribution of the total, mapped and unmapped reads were similar. Moreover, the number of mapped reads was not dependent on the length of genomic contigs, but reflected the abundance of particular reads in the transcriptome (p<0.05). For example, whereas 265 reads mapped onto H. glycines genomic contig ABLA01000003.1 (21602 nt), only 56 mapped to contig ABLA01000001.1 (24,040 nt). On average 9 reads mapped to the H. glycines contigs with a range of 1 to 5,901 and with an average read consensus length of 603 nt. The read mapping could clearly be used to delineate intron-exon boundaries of the genomic sequences. For example, H. glycines genomic contig with the longest matched read consensus, contig ABLA01000003, contains genes putatively encoding four C. elegans homologues namely dynein heavy chains (dhc-1, che-3), ribosomal protein subunit 31 (rpl-31) and deoxyuridine triphosphate nucleotidohydrolase (dut-1).
A total of 454 out of the 458 Core Eukaryotic Genes (CEGs) were identified altogether in the J2 transcriptome of H. schachtii and in the genomic contigs of the four PPNs: of these 256 were common. For the H. schachtii transcriptome, about 2% of transcripts (1149 contigs and 905 singletons) were similar to a total of 419 CEGs. Varying numbers of genomic contigs of the four PPNs identified from BLAST comparisons appear to encode CEGs; 15.3% for M. hapla, 12% for M. incognita, 3.1% for H. glycines and 5.2% for G. pallida (Fig 5C). Relatively more CEGs were identified in the transcriptome and the genome of H. glycines than the other genomes and these two species also had more CEGs in common (Fig 5C).
The spliceosome of H. schachtii: evidence of trans-splicing
Very little is known about the components/genes of the spliceosome and the mechanism of trans-splicing in PPNs. We used the 104 genes involved in the KEGG spliceosome pathway of C. elegans to identify orthologues in the H. schachtii transcriptome and compared these to those of the PPNs H. glycines, M. incognita, M. hapla, P. thornei, P. coffeae, G. rostochiensis and G. pallida. The results are presented as a heat map with colours indicating relative values of the bit score from the alignments (Fig 6). Using a threshold bit score of 100, twenty of the 104 genes were present in all the PPNs with prp-8 (pre-mRNA-processing factor 8), the most conserved. In all, 56 homologues were identified in the H. schachtii transcriptome compared to 81 in ESTs of H. glycines.
Global analysis shows that over 70% of C. elegans expressed genes are trans-spliced to one of two 22 nucleotide splice leaders [35]. Using the regular expression derived from the conserved regions of SL1 RNA genes of C. elegans and P. pacificus, which usually has the canonical 22 nt SL1 leader sequence (GGTTTAATTACCCAAGTTTGAG) and a Sm binding site (ATTTTGGAAC)[32], we searched for existence of a similar SL1 RNA gene in the genomic sequences of H. glycines using BLASTN and manual curation. Of the 45,526 genomic contigs of H. glycines deposited at NCBI, 218 had at least one variant (with 0–3 mismatches) of the SL1 sequence above: for 182 of these the SL1 was a 100% match to the query, 23 others had the maximum of three mismatches and 13 with one or two mismatches. Genomic contig ABLA01003722.1 contained the maximum number of five SL1s, approximately 600 bases apart at positions 541, 1161, 1770, 2380 and 2990. Of the sequences with SL1, 152 also had a sequence similar to the Sm binding site. Using this information, we identified 823 H. schachtii reads with variants of the SL1 sequence. There were 587 reads with 100% match to the SL1 sequence. The reads ranged from 42 to 561 nt long with only 75 of them <100 nt long. About 260 of the transcripts were annotated (e-value of 1E-05). Of these, 71 putatively encoded ribosomal proteins and subunits including rps-7, rps-15, rps-23 and rpl-19 homologues of C. elegans, G. rostochiensis, G. pallida, B. xylophilus, B. malayi and L. loa. Others were similar to transcripts for vacuolar H ATPase protein 16 (vha-16), 1, 4-beta endoglucanase, splicing factors (e.g. rsp-6), FMRFamide-like protein 5 and 16 (flp-5, flp-16), expansin B1, troponin C (pat-10), synaptobrevin (snb-1), mucin-19-like protein, heat shock factor-binding protein 1-like and cold-shock-like protein (csp-1). None of the SL2-like sequences found in the Rhabditina familiy was associated with any transcript.
Transcripts putatively encoding carbohydrate active enzymes
Using BLASTX to search the 188,623 protein sequences of the CAZy database, a total of 407 H. schachtii transcripts (226 contigs and 182 singletons) matched sequences of Glycoside Hydrolases (GHs), Carbohydrate Binding Modules (CBMs), Polysaccharide Lyases (PLs), Carbohydrate Esterases (37), Glycosyl Transferases (GTs) and proteins of Auxiliary Activities (AA) of other organisms. Of these, 375 had a maximum HSP score of at least 50 to the best matching CAZyme (S3 Table). The matching CAZymes were from the following CAZy families: eight CBMs, five AAs, three CEs, 39 GHs, two PLs and 31 GTs and were mostly sequences of other nematodes particularly Caenorhabditis spp., A. suum, Strongyloides ratti and plant parasitic nematodes.
The number of transcripts (36) matching to CAZymes of plant parasitic nematodes is shown in Table 1. Five contigs and four singletons matched PL3 pectate lyases of B. xylophilus (AGI04333.1), Globodera tabacum tabacum (AEA08812.1), H. glycines (AAM74954.1, ADW77535.1, ADW77534.1), and a pectate lyase precursor of H. schachtii (ABN14273.1, ABN14272.1). Fourteen contigs and five singletons putatively encode GH5 CAZymes as these transcripts had high percentage similarities to those of enzymes identified for other plant parasitic nematodes of six genera including those characterised for H. schachtii and the closely related H. glycines (Table 1). Twenty five transcripts matched GH CAZymes with multiple domains and those from PPNs were usually GH5s with CBM2 family (Table 1 and S3 Table). Together with pectate lyases which act on pectin components of plant cell walls, GH5 cellulases or beta 1,4 endoglucanases, secreted from gland cells, are generally characterised in most PPNs and are thought to modify cell walls, mainly facilitating intracellular migration of nematode juveniles through host tissues but possibly also involved in syncytial expansion [7]. In addition, two contigs matched a GH53 arabinogalactan endo-1,4-beta-galactosidase 1 of H. schachtii, an enzyme capable of hydrolysing beta-1,4-galactan: such a protein could play a role in modifying plant cell wall during migration of the nematode through roots [36].
A majority (21) of the best CAZyme matches from the 31 GT families were to sequences of Caenorhabditis spp. and A. suum and were mostly enzymes involved in common molecular processes of development, for example N-acetyllactosamine synthase, glycogen phosphorylase and chitin synthase (S3 Table). The only CAZyme of a plant parasitic nematode origin with a GT activity was putative trehalose 6-phosphate synthase of Aphelenchus avenae; this enzyme is involved in metabolism of trehalose which is important in several developmental processes including energy reservation, egg hatching and protection from biotic stresses [37]. The best CAZyme matches to the H. schachtii transcripts for some families were of non-nematode origins. These were sequences of plant, fungi, bacteria and insect species (S3 Table).
To confirm that these were not contaminants in the transcriptome, the matching CAZymes were compared to nematode sequences in the NCBI databases using BLAST. All but eight of the 40 CAZymes matched significantly to sequences of various nematode species including Caenorhabditis spp, S. ratti, Trichinella spp and plant parasitic nematodes. The presence of transcripts matching the eight CAZymes with no significant match to any other nematode sequence in the NCBI database needs further investigation.
H. schachtii orthologues of putative nematode parasitism genes
Plant parasitic nematodes successfully parasitise their hosts by employing secretions from pharyngeal gland cells, amphids and the hypodermis: the secreted peptides, which are responsible for various activities, are generally described as ‘effectors’. These are thought to play a number of roles which include suppression of host defences, enabling migration in plant tissues, promotion of nematode feeding, formation of feeding tubes, digestion of ingested cytoplasm, and host cell modification leading to the induction and maintenance of feeding structures such as syncytia for cyst nematodes or giant cells for root knot nematodes. To assess whether the pre-infective J2s of H. schachtii are primed for infection, we compared the transcriptome to sequences of 30 genes (29 complete cDNAs and one partial cDNA) representative of the effector repertoire of plant parasitic nematodes: for each of these there is evidence of secretion from gland cells and/or an effector function during interaction with host plants.
In all, 148 contigs and 183 singletons were identified with sequence similarity to 25 of the effectors (Table 2). These included proteins that can modify host metabolic profiles (e.g. chorismate mutase), and those that may aid digestion of host cell contents (cathepsin L) or are potentially secreted to protect the nematode against host defence (e.g. peroxiredoxin). Also there were transcripts with similarity to MAP-1 gene of Meloidogyne species, which is potentially involved in the early stages of host recognition [38]. Transcripts of high sequence identity to the Hg30C02 effector of H. glycines, expression of which increases susceptibility of Arabidopsis to infection by H. schachtii and the Hs19C07 effector, which interacts with the Arabidopsis auxin influx transporter LAX3 to facilitate syncytium development, were identified [5, 39, 40]. There were two transcripts with low HSP scores to the 10A06 effector of H. schachtii: expression of this gene induces morphological changes in the host, targets spermidine synthase and possibly disrupts Arabidopsis defense signalling leading to increased susceptibility to infection [41].
In addition to transcripts putatively encoding the cell wall-modifying CAZymes, beta 1, 4 endoglucanase, pectate lyase and arabinogalactan endo-1, 4-beta-galactosidase 1, H. schachtii transcripts with relatively low sequence similarity to two other cell wall-modifying enzymes were identified from the transcriptome; one transcript for polygalacturonase (M. incognita) and four transcripts for expansin (G. rostochiensis) (Table 1). However, no transcripts were found with sequence similarity to those of cell wall-modifying enzymes beta-1,3-endoglucanase (B. xylophilus), GHF45 family protein (B. xylophilus) and xylanase (xyl-1) (M. incognita), and none matched gene sequences of cellulose binding protein precursor (cbp-1) of M. javanica and a chitinase (H. glycines). The latter has not been characterised in detail, but accumulates specifically in the subventral oesophageal gland cells of parasitic stages of H. glycines, although not in eggs or hatched pre-parasitic second-stage juveniles (Table 2) [42].
Effectors secreted by the hypodermis and amphids of plant parasitic nematodes have not been well-studied. In this study, we identified transcripts that potentially encode similar putative effectors secreted from the amphid (e.g. Ams1) and hypodermis (e.g. sxp1) of G. rostochiensis (Table 2). Recently, amphid-secreted effectors of G. pallida, HYP effectors, that have hyper-variable regions in individuals of the nematode population have been characterised [65]. No H. schachtii transcript with significant identity to sequences of these effectors was identified.
Various EST libraries of gland cells and contents have been constructed for the cyst nematode H. glycines and the root knot nematode M. incognita and are available in the NCBI databases [9, 66–69]. Whilst a small number of these transcripts have been studied in detail, a majority remain uncharacterised. A total of 11,561 H. schachtii transcripts matched 4,995 of the 15,226 gland cell-derived ESTs of H. glycines mostly at e-values much lower than the 1E-05 threshold and with very high HSP scores, the highest at 597 with an average of 93. Of the 40 putative oesophageal gland cell secretory proteins (msp1-40) for M. incognita [67], six (msp10, 21, 26, 28, 29, 34 and 40) had matches to a total of 35 H. schachtii transcripts with e-values between 1E-05 and 1E-14 and maximum HSP scores between 31 and 69. Generally, matching H. schachtii transcripts had high identity to (putative) effectors of cyst nematodes compared to those of root knot nematodes. When SignalP4.1 was used to predict the presence and location of signal peptide cleavage sites putatively encoded by all transcripts with matches to those of nematode effectors and CAZymes, four fully translated transcripts putatively encoding >50 amino acids were identified (S4 Table). These transcripts were most similar to those of three effectors of H. glycines (annexin 4C10, chorismate mutase, the gland protein G19C07) and calreticulin of M. incognita. None of the four transcripts was predicted to have a transmembrane helix; this feature and the presence of a signal peptide are two important features of nematode effectors that are secreted and transported into host cells.
RNAi genes identified in the H. schachtii transcriptome
RNAi is a natural gene regulation mechanism in eukaryotes. Genes involved in RNAi pathways in C. elegans are well-characterised and have been used as a model to understand the process in other organisms. It is known that different numbers of genes are involved in the various processes of this otherwise conserved process in different organisms [70]. Moreover, in PPNs there seem to be differences in susceptibility to RNAi between cyst and root knot nematodes, and also within Pratylenchus species [24]. The recent sequencing of genomes of M. incognita, M. hapla and G. pallida provides information on RNAi genes in these organisms. However, there is currently no information on similar genes for H. schachtii. To identify such genes from the transcriptome, we compared the transcripts to protein sequences of 97 functionally characterised RNAi effectors of C. elegans (S1 Text) including 27 argonautes and to those identified in published transcriptomes of P. coffeae, H. avenae and the genomes of M. incognita, M. hapla and G. pallida.
There were significant matches to 36 of the 97 RNAi effectors characterised for C. elegans (Table 3). Interestingly, at least one of the genes was involved in all the major functional classes of the RNAi machinery i.e. spreading of RNAi triggers into and out of cells (e.g. xpo-1), the dicer complex (e.g. dcr-1, drh-1), the RNAi-Inducing Silencing Complex (e.g. ain-1), RNAi amplification (e.g. ego-1), RNAi inhibitors (e.g. eri-1) and nuclear RNAi effectors (cid-1). In addition, transcripts similar to two prominent genes involved in the processing of primary microRNA transcripts, drsh-1 and pash-1, and for argonautes including worm-specific argonautes (WAGOs) were also identified in the transcriptome. More genes putatively encoding RNAi effectors were identified in this study than the number published for transcriptomes of P. coffeae and H. avenae and genomes of M. incognita and G. pallida [15, 17, 19, 71](Table 3). A possible explanation for this difference is the larger pool of C. elegans RNAi effectors used for this study, which included those recently characterised. Transcripts for only seven of the RNAi effectors were identified in all five nematodes. With the exception of the P. coffeae transcriptome, putative RNAi effectors for each of the functional classes of RNAi had been identified (Table 3). This information provides a template for comprehensive study of RNAi in cyst nematodes and how the phenomenon can be applied as a potential strategy to control PPNs.
H. schachtii may harbour virus-like genomes
A recent analysis showed that the transcriptome of H. glycines was associated with virus-like genomes, and genome fragments of four different RNA viruses were found in the transcriptome, including the Soybean cyst nematode nyavirus (ScNV) [23]. Using TBLASTX, 278 of the transcriptome reads were similar to the full-length sequence of the Soybean cyst nematode nyavirus (ScNV, PRJNA258186), the best was 461 nt long (e-value 7.24E-80, bit score 1,366). This similarity was confirmed using read mapping where 248 reads were mapped to the genome. Thirty nine contigs of the H. schachtii J2 transcriptome had high identities to the ScNV genome, the best contig with e-value of 2E-175, total bit score of 1,373 and maximum HSP score of 534. The contig with the best alignment matched the putative RNA dependent RNA polymerase protein (AEF56729.1). This contained an open reading frame of 663 amino acids and matched the viral protein with a maximum score of 885 and with 63% protein identity. Also, two contigs matched the glycoprotein (AEF56728.1) with an e value of 9E-114 and a bit score of 372. Transcripts with identity to two conserved domains characteristic of RNA dependent RNA polymerase of members of the Mononegavirales, matched the phosphoprotein, P (grey residues in Fig 7), which acts as a transcription factor and the large protein L, which confers the RNA polymerase activity on the complex and carries a motif, GxxTx(n)HR, that is essential for mRNA cap formation. Notably, no H. schachtii read or contig matched the ORF2 and ORF3 that encode the hypothetical proteins of the ScNV.
Discussion
The J2 stage of a cyst nematode is often viewed as the most important stage in its life cycle because this life stage must be able to move through soil to locate a host root, enter and migrate within it while evading or suppressing the host immune responses, to a site where it initiates the formation of a syncytium. The J2 stage can be divided into pre- and post-plant entry stages. In addition to expression of genes involved in general metabolism, the results presented here clearly show that even before root entry the pre-parasitic J2 is already primed to express many parasitism-related genes of varied functions needed for it to reach host roots, enter them and migrate through them, evade host defences, and initiate syncytium development. Post-root entry J2s may well express additional parasitism genes, and there will undoubtedly be changes in the levels of many transcripts as the nematodes become sedentary, expand and reproduce as has been found for G. pallida [72]. A more detailed stage-specific set of transcriptome data is needed to follow changes in gene expression for post-entry J2s and subsequent developmental stages. However, the data generated here are sufficient to identify sequences of potential target genes that can be used to develop new methods to control pre-parasitic J2s, and so reduce or prevent root infestation.
The analysis of the pre-parasitic J2 transcriptome of H. schachtii has thus enabled us to identify genes expressed at this stage and with possible roles in host parasitism. Many of the transcripts were similar to genes found in other endoparasitic nematodes, whilst some were exclusive to cyst nematodes. Not surprisingly more of the transcripts share sequence homology to those of cyst nematodes with about 67% of the reads mapping to genomic sequences of the closely related H. glycines. Transcripts similar to those of 2,410 C. elegans proteins were identified including genes involved in splicing and trans-splicing and those with direct roles in RNA interference. Some of those transcripts with no homology to genes of free-living nematodes were identified as common to PPNs and may well be required for parasitism in general or specifically by cyst nematodes for plant parasitism.
Once inside the root J2s of H. schachtii typically migrate intracellularly through cells to a site at which they establish a metabolically active syncytium [10, 12]. They need to secrete enzymes that can modify key polysaccharide components of host cell walls (cellulose, hemicelluloses and pectin) to facilitate these activities. The expression before root entry of a range of such transcripts encoding carbohydrate active enzymes clearly shows that pre-parasitic J2s of H. schachtii are primed for host infection. Examples include transcripts of the beta-1,4- endoglucanase and its precursors, a cellulase with GH5 enzyme activity which catalyses the hydrolysis of the glycosidic bonds in cellulose. Similar transcripts for cellulases found here had high percentage sequence identity to those previously characterised for H. schachtii, and two other cyst nematodes, G. tabacum solanacearum and H. glycines. Some were predicted to have signal peptides and carbohydrate active binding modules, both of which are characteristics of cellulases widely deployed by PPNs including migratory endoparasites such as P. thornei [20]. The identification of transcripts of pectate lyase, which is involved in modification of pectic polymers, is consistent with the expression and deployment of this gene product by other cyst nematodes during infection [13].
Consistent with other studies, for most effectors of root knot nematodes there was either no similar transcript in the H. schachtii transcriptome or matching transcripts had very low similarity scores. These genes may be absent from the genome of H. schachtii or they may be present but not expressed during early parasitism. To validate this observation, four characterised effectors of M. incognita, xylanase (MiXly, AF224342), NodL factor (AW829666.1), polygalacturonase (AY098646.1), and 16D10 (Q6YKB1.1) were used as references to identify similar transcripts from the genomic sequences of the cyst nematodes H. glycines and G. pallida using BLASTX at an e-value cut-off 1E-03. As with the H. schachtii transcripts, no genomic contig of either cyst nematode genome appears to encode any protein similar to Mixyl and Mi16D10. For NodL, whilst there was no match to any H. schachtii transcript, five genomic contigs of H. glycines (the best with a maximum bit score of 131) and one for G. pallida with low maximum bit score (34) were identified. There was only one contig each from the transcriptome of H. schachtii (at e-value 1.09E-5) and genome of G. pallida (at e-value 9.33E-5), but none from H. glycines that appears to encode a polygalacturonase similar to that of root knot nematodes. Differences in the complements of effectors, particularly cell wall modifying enzymes, probably reflect the different modes of migration through host roots: J2 root knot nematodes migrate intercellularly through cells walls rather than intracellularly from cell to cell, and so require a different set of enzymes to modify cell walls: fewer such enzymes would be needed for intracellular migration by J2 cyst nematodes [21, 73].
In addition to intracellular migration in roots, endoparasitic J2 and subsequent stages must be able to evade or modulate the host immune system. Some effectors involved in these activities may be secreted via the stylet, but others thought to be involved in this process may be synthesised and secreted directly via the amphids, or via the hypodermis after being transported across the cuticle, which is in direct contact with the external environment. Such effectors include the MAP-1 in J2 amphidial secretions of M. incognita possibly involved in the early stages of recognition between resistant plants and avirulent nematodes [38], and enzymes known to regulate plant defense responses or signalling such as peroxiredoxin (Gr-tpx), glutathione peroxidase (Gr-gpx1), a glutathione S-Transferase (Mi-gst-1) which counter reactive oxygen species mediated signalling, and a fatty acid and retinol binding protein (FAR-1, G. pallida, M. incognita) present on cuticular surfaces of nematodes. The latter appear to bind lipid precursors of plant defence compounds and prevent their metabolism [44, 74]. Transcripts with significant similarity to two putative effectors of G. rostochiensis, Gr-ams1 and Gr-sxp-1, secreted from the amphids and hypodermis respectively, were also identified. Another class of hyper-variable apoplastic effector gene family termed HYP has been characterised in G. pallida and is apparently only present in cyst nematodes and Rotylenchulus reniformis. HYP effectors are secreted from the amphids, and are required for successful infection. Interestingly, full length transcripts identical to these effectors were not found in the pre-J2 H. schachtii transcriptome [65]. The identification of transcripts encoding similar effectors including the SXP-RAL2 gene family, secreted from the epidermis and the amphidial sheath cells in G. rostochiensis [45] indicate these effectors are synthesised in the pre-parasitic J2, pre-entry and not necessarily synthesised in response to plant defence signalling.
Two well-characterised secreted proteins of cyst nematodes are known to be functionally similar to and can mimic the functions of host plant orthologues [50, 75]. These are the CLE-like proteins and chorismate mutase: transcripts of both were present in the pre-parasitic J2 transcriptome. Chorismate mutases of plant parasites (e.g. nematodes and fungi) change the metabolic status of host cells through metabolic priming or alter the synthesis of chorismate-derived products, resulting in a down-regulation of a host plant defense [50, 76–78]. Similar transcripts for the two proteins and for those of several secreted proteins have been shown to express in the single dorsal and/or the two subventral oesophageal cells of pre-parasitic and parasitic stages of H. glycines and G. rostochiensis [9, 78, 79]. Complete sequences of orthologues of most of these genes have not been found in the transcriptomes of migratory endoparasites [19–21]. It is possible such genes are required specifically by cyst nematodes to initiate or maintain feeding sites by modulating plant growth or metabolic processes at feeding sites, or there may be orthologues present in migratory nematodes that have yet to be characterised.
There are reports that both free-living (C. elegans and C. briggsae) and some PPNs (e.g. H. glycines) can harbour viruses [23, 80]. In the transcriptome of J2 H. schachtii reads mapped to the genome sequence of ScNV and there was a significant match of contigs to three of the five open reading frames of this virus: this provides good evidence that a virus similar to ScNV is also present in H. schachtii. New sequencing technologies have revealed the presence of many new viruses in animals and plants, and have given rise to a view that not all viruses are pathogens, some are vertically transmitted, and some can be mutualistic or beneficial [81]. The source of viruses in PPN genomes is not known at present; possible sources are the plants from which they feed or from the soil environment. J2s of sedentary nematodes, which are obligate parasites, usually find a host to feed from soon after hatching. In the case of the H. schachtii analysed here, because the J2s had not fed from host plants, the source of infection is either from soil or was acquired through vertical transmission from the preceding generation. Functional characterisation of these sequences and other transcripts identical to uncharacterised gene products could provide additional insight into the parasitic life of H. schachtii and potentially to other parasitic nematodes. Some of the transcripts may be involved in development and metabolism of the nematode, but others could be involved in nematode-host interactions.
In this work we have demonstrated that the pre-parasitic J2 stage of H. schachtii is primed for host root entry, and expresses many of the transcripts whose products will be employed in root entry, migration through host tissues, evasion or modification of host defences, and also some of the effectors thought to be required for induction of syncytial feeding cells. The identification of a substantial set of candidate parasitism genes also provides some new targets for their control. Evidence is also provided for the presence of viral sequences in J2s of H. schachtii: this aspect and the presence of many other unannotated transcripts require further study, for example using RNAi, which has been used to characterise some effectors found in this study.
Supporting Information
S1 Fig. A mist apparatus for extracting nematodes from soil.
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S1 Table. Sequences and databases used for annotation of the H. schachtii transcriptome.
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S2 Table. PAMGO terms associated with H. schachtii transcripts.
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S3 Table. H. schachtii transcripts putatively encoding carbohydrate active enzymes.
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S4 Table. H. schachtii transcripts predicted to encode signal peptides without trans-membrane helices.
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S1 Text. Translated sequences of RNAi genes of C. elegans used for identifying similar transcripts of H. schachtii.
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Acknowledgments
PN was supported by a PhD studentship from Murdoch University; RG was supported by an Australian India Strategic Research Fund Grant (BF030027); Nemgenix Pty. Ltd. supported JFN and contributed funds for consumables. We thank members of the Plant Biotechnology Research Group (made up of Academics, Post-Doctoral researchers and students) at Murdoch University for meaningful discussions.
Author Contributions
Conceived and designed the experiments: JFN MGKJ. Performed the experiments: JFN RG. Analyzed the data: PN JFN FN. Contributed reagents/materials/analysis tools: JFN MGKJ. Wrote the paper: PN JFN MGKJ.
References
- 1. Abawi GM, Mai WF. Effects of initial population densities of Heterodera schachtii on yield of cabbage and table beets in New York State. Phytopathology. 1980;70(6):481–5.
- 2. Muller J. The economic importance of Heterodera schachtii in Europe. Helminthologia. 1999;36(3):205–13.
- 3. Burt OR, Ferris H. Sequential decision rules for managing nematodes with crop rotations. J Nematol. 1996;28(4):457–74. PMC2619718. pmid:19277164
- 4. Fosu-Nyarko J, Jones MG. Chapter Fourteen-Application of biotechnology for nematode control in crop plants. Adv Bot Res. 2015;73:339–76.
- 5. Fuller VL, Lilley CJ, Urwin PE. Nematode resistance. New Phytol. 2008;180(1):27–44. pmid:18564304
- 6. Sijmons PC, Grundler FMW, von Mende N, Burrows PR, Wyss U. Arabidopsis thaliana as a new model host for plant-parasitic nematodes. Plant J. 1991;1(2):245–54.
- 7. Smant G, Stokkermans JPWG, Yan Y, de Boer JM, Baum TJ, Wang X, et al. Endogenous cellulases in animals: Isolation of β-1,4-endoglucanase genes from two species of plant-parasitic cyst nematodes. Proc Natl Acad Sci USA. 1998;95(9):4906–11. pmid:9560201
- 8. Vanholme B, Mitreva M, Van Criekinge W, Logghe M, Bird D, McCarter J, et al. Detection of putative secreted proteins in the plant-parasitic nematode Heterodera schachtii. Parasitol Res. 2006;98(5):414–24. pmid:16380840
- 9. Gao B, Allen R, Maier T, Davis EL, Baum TJ, Hussey RS. The parasitome of the phytonematode Heterodera glycines. Mol Plant-Microbe In. 2003;16(8):720–6.
- 10. Jones MGK. Host cell responses to endoparasitic nematode attack: structure and function of giant cells and syncytia. Ann Appl Biol. 1981;97(3):353–72.
- 11. Jones MGK, Dropkin VH. Cellular alterations induced in soybean roots by three endoparasitic nematodes. Physiol Plant Pathol. 1975;5(2):119–24.
- 12. Jones MGK, Northcote DH. Nematode-induced syncytium-a multinucleate transfer cell. J Cell Sci. 1972;10:789–809. pmid:5038416
- 13. Vanholme B, Van Thuyne W, Vanhouteghem K, De Meutter JAN, Cannoot B, Gheysen G. Molecular characterization and functional importance of pectate lyase secreted by the cyst nematode Heterodera schachtii. Mol Plant Pathol. 2007;8(3):267–78. pmid:20507498
- 14. Hewezi T, Howe P, Maier TR, Hussey RS, Mitchum MG, Davis EL, et al. Cellulose binding protein from the parasitic nematode Heterodera schachtii interacts with Arabidopsis pectin methylesterase: cooperative cell wall modification during parasitism. Plant Cell. 2008;20(11):3080–93. pmid:19001564
- 15. Abad P, Gouzy J, Aury J- M, Castagnone-Sereno P, Danchin EGJ, Deleury E, et al. Genome sequence of the metazoan plant-parasitic nematode Meloidogyne incognita. Nat Biotechnol. 2008;26(8):909–15. http://www.nature.com/nbt/journal/v26/n8/suppinfo/nbt.1482_S1.html. pmid:18660804
- 16. Opperman CH, Bird DM, Williamson VM, Rokhsar DS, Burke M, Cohn J, et al. Sequence and genetic map of Meloidogyne hapla: A compact nematode genome for plant parasitism. Proc Natl Acad Sci USA. 2008;105(39):14802–7. pmid:18809916
- 17. Cotton J, Lilley C, Jones L, Kikuchi T, Reid A, Thorpe P, et al. The genome and life-stage specific transcriptomes of Globodera pallida elucidate key aspects of plant parasitism by a cyst nematode. Genome Biol. 2014;15(3):R43. pmid:24580726
- 18. Burke M, Scholl EH, Bird DM, Schaff JE, Coleman S, Crowell R, et al. The plant parasite Pratylenchus coffeae carries a minimal nematode genome. Nematology. 2015;17(6):621–37.
- 19. Haegeman A, Joseph S, Gheysen G. Analysis of the transcriptome of the root lesion nematode Pratylenchus coffeae generated by 454 sequencing technology. Mol Biochem Parasit. 2011;178(1–2):7–14.
- 20. Nicol P, Gill R, Fosu-Nyarko J, Jones MGK. de novo analysis and functional classification of the transcriptome of the root lesion nematode, Pratylenchus thornei, after 454 GS FLX sequencing. Int J Parasitol. 2012;42(3):225–37. pmid:22309969
- 21. Jones MGK, Fosu-Nyarko J. Molecular biology of root lesion nematodes (Pratylenchus spp.) and their interaction with host plants. Ann Appl Biol. 2014;164(2):163–81.
- 22. Fosu-Nyarko J, Tan J- ACH, Gill R, Agrez VG, Rao U, Jones MGK. De novo analysis of the transcriptome of Pratylenchus zeae to identify transcripts for proteins required for structural integrity, sensation, locomotion and parasitism. Mol Plant Pathol. 2015:
- 23. Bekal S, Domier LL, Niblack TL, Lambert KN. Discovery and initial analysis of novel viral genomes in the soybean cyst nematode. J Gen Virol. 2011;92(8):1870–9.
- 24. Tan J-ACH, Jones MGK, Fosu-Nyarko J. Gene silencing in root lesion nematodes (Pratylenchus spp.) significantly reduces reproduction in a plant host. Ex Parasitol. 2013;133(2):166–78.
- 25. Huang X, Madan A. CAP3: A DNA Sequence Assembly Program. Genome Res. 1999;9(9):868–77. pmid:10508846
- 26. Altschul SF, Madden TL, Schäffer AA, Zhang J, Zhang Z, Miller W, et al. Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 1997;25(17):3389–402. pmid:9254694
- 27. Hall TA. BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucl Acids Symp Ser. 1999;41:95–8.
- 28. Consortium GO. The Gene Ontology (GO) project in 2006. Nucleic Acids Res. 2006;34(suppl 1):D322–D6.
- 29. Cantarel BL, Coutinho PM, Rancurel C, Bernard T, Lombard V, Henrissat B. The Carbohydrate-Active EnZymes database (CAZy): an expert resource for Glycogenomics. Nucleic Acids Res. 2009;37(suppl 1):D233–D8.
- 30. Torto-Alalibo T, Collmer C, Gwinn-Giglio M. The Plant-Associated Microbe Gene Ontology (PAMGO) Consortium: community development of new Gene Ontology terms describing biological processes involved in microbe-host interactions. BMC Microbiology. 2009;9(Suppl 1):S1. pmid:19278549
- 31. Parra G, Bradnam K, Korf I. CEGMA: a pipeline to accurately annotate core genes in eukaryotic genomes. Bioinformatics. 2007;23(9):1061–7. pmid:17332020
- 32. Lee K-Z, Sommer RJ. Operon Structure and trans-splicing in the nematode Pristionchus pacificus. Mol Biol Evol. 2003;20(12):2097–103. pmid:12949121
- 33. Guiliano DB, Blaxter ML. Operon conservation and the evolution of trans-splicing in the Phylum Nematoda. PLoS Genet. 2006;2(11):e198. pmid:17121468
- 34. Petersen TN, Brunak S, von Heijne G, Nielsen H. SignalP 4.0: discriminating signal peptides from transmembrane regions. Nat Methods. 2011;8(10):785–6. pmid:21959131
- 35. Allen MA, Hillier LW, Waterston RH, Blumenthal T. A global analysis of C. elegans trans-splicing. Genome Res. 2011;21(2):255–64. pmid:21177958
- 36. Vanholme B, Haegeman A, Jacob J, Cannoot B, Gheysen G. Arabinogalactan endo-1, 4-β-galactosidase: a putative plant cell wall-degrading enzyme of plant-parasitic nematodes. Nematology. 2009;11(5):739–47.
- 37. Behm CA. The role of trehalose in the physiology of nematodes. Int J Parasitol. 1997;27(2):215–29. pmid:9088992
- 38. Semblat J-P, Rosso M-N, Hussey RS, Abad P, Castagnone-Sereno P. Molecular cloning of a cDNA encoding an amphid-secreted putative avirulence protein from the root-knot nematode Meloidogyne incognita. Mol Plant-Microbe In. 2001;14(1):72–9.
- 39. Lee C, Chronis D, Kenning C, Peret B, Hewezi T, Davis EL, et al. The novel cyst nematode effector protein 19C07 interacts with the Arabidopsis auxin influx transporter LAX3 to control feeding site development. Plant Physiol. 2011;155:866–80. pmid:21156858
- 40. Hamamouch N, Li C, Hewezi T, Baum TJ, Mitchum MG, Hussey RS, et al. The interaction of the novel 30C02 cyst nematode effector protein with a plant β-1, 3-endoglucanase may suppress host defence to promote parasitism. J Exp Bot. 2012;63(10):3683–95. pmid:22442414
- 41. Hewezi T, Howe PJ, Maier TR, Hussey RS, Mitchum MG, Davis EL, et al. Arabidopsis spermidine synthase is targeted by an effector protein of the cyst nematode Heterodera schachtii. Plant Physiol. 2010;152(2):968–84. pmid:19965964
- 42. Gao B, Allen R, Maier T, McDermott JP, Davis EL, Baum TJ, et al. Characterisation and developmental expression of a chitinase gene in Heterodera glycines. Int J Parasitol. 2002;32(10):1293–300. pmid:12204229
- 43. Dubreuil G, Magliano M, Deleury E, Abad P, Rosso M. Transcriptome analysis of root—knot nematode functions induced in the early stages of parasitism. New Phytol. 2007;176(2):426–36. pmid:17692078
- 44. Prior A, Jones J, Blok V, Beauchamp J, Mcdermott L, Cooper A, et al. A surface-associated retinol-and fatty acid-binding protein (Gp-FAR-1) from the potato cyst nematode Globodera pallida: lipid binding activities, structural analysis and expression pattern. Biochem J. 2001;356:387–94. pmid:11368765
- 45. Jones JT, Smant G, Blok VC. SXP/RAL-2 proteins of the potato cyst nematode Globodera rostochiensis: secreted proteins of the hypodermis and amphids. Nematology. 2000;2(8):887–93.
- 46. Lozano-Torres JL, Wilbers RH, Gawronski P, Boshoven JC, Finkers-Tomczak A, Cordewener JH, et al. Dual disease resistance mediated by the immune receptor Cf-2 in tomato requires a common virulence target of a fungus and a nematode. Proc Natl Acad Sci USA. 2012;109(25):10119–24. pmid:22675118
- 47. Jones J, Reavy B, Smant G, Prior A. Glutathione peroxidases of the potato cyst nematode Globodera rostochiensis. Gene. 2004;324:47–54. pmid:14693370
- 48. Jones JT, Kumar A, Pylypenko LA, Thirugnanasambandam A, Castelli L, Chapman S, et al. Identification and functional characterization of effectors in expressed sequence tags from various life cycle stages of the potato cyst nematode Globodera pallida. Mol Plant Pathol. 2009;10(6):815–28. pmid:19849787
- 49. Qin L, Kudla U, Roze EH, Goverse A, Popeijus H, Nieuwland J, et al. Plant degradation: a nematode expansin acting on plants. Nature. 2004;427(6969):30-. pmid:14702076
- 50. Bekal S, Niblack TL, Lambert KN. A chorismate mutase from the soybean cyst nematode Heterodera glycines shows polymorphisms that correlate with virulence. Mol Plant-Microbe In. 2003;16(5):439–46.
- 51. Carpentier J, Esquibet M, Fouville D, Manzanares-Dauleux MJ, Kerlan MC, Grenier E. The evolution of the Gp—Rbp—1 gene in Globodera pallida includes multiple selective replacements. Mol Plant Pathol. 2012;13(6):546–55. pmid:22192092
- 52. Postma WJ, Slootweg EJ, Rehman S, Finkers-Tomczak A, Tytgat TO, van Gelderen K, et al. The effector SPRYSEC-19 of Globodera rostochiensis suppresses CC-NB-LRR-mediated disease resistance in plants. Plant Physiol. 2012;160(2):944–54. pmid:22904163
- 53. Ali S, Magne M, Chen S, Côté O, Stare BG, Obradovic N, et al. Analysis of putative apoplastic effectors from the nematode, Globodera rostochiensis, and Identification of an expansin-like protein that can induce and suppress host defenses. PloS one. 2015;10(1):e0115042. pmid:25606855
- 54. Robertson L, Robertson WM, Sobczak M, Helder J, Tetaud E, Ariyanayagam MR, et al. Cloning, expression and functional characterisation of a peroxiredoxin from the potato cyst nematode Globoderarostochiensis. Mol Biochem Parasit. 2000;111(1):41–9.
- 55. Neveu C, Abad P, Castagnone-Sereno P. Molecular cloning and characterization of an intestinal cathepsin L protease from the plant-parasitic nematode Meloidogyne incognita. Physiol Mol Plant P. 2003;63(3):159–65.
- 56. Jaouannet M, Perfus—Barbeoch L, Deleury E, Magliano M, Engler G, Vieira P, et al. A root—knot nematode—secreted protein is injected into giant cells and targeted to the nuclei. New Phytol. 2012;194(4):924–31. pmid:22540860
- 57. Jaubert S, Laffaire J-B, Ledger T, Escoubas P, Amri E-Z, Abad P, et al. Comparative analysis of two 14-3-3 homologues and their expression pattern in the root-knot nematode Meloidogyne incognita. Int J Parasitol. 2004;34(7):873–80. pmid:15157770
- 58. Jaouannet M, Magliano M, Arguel M, Gourgues M, Evangelisti E, Abad P, et al. The root-knot nematode calreticulin Mi-CRT is a key effector in plant defense suppression. Mol Plant-Microbe In. 2013;26(1):97–105.
- 59. Jaubert S, Laffaire J-B, Abad P, Rosso M-N. A polygalacturonase of animal origin isolated from the root-knot nematode Meloidogyne incognita. FEBS letters. 2002;522(1):109–12.
- 60. Jacob J, Vanholme B, Haegeman A, Gheysen G. Four transthyretin-like genes of the migratory plant-parasitic nematode Radopholus similis: members of an extensive nematode-specific family. Gene. 2007;402(1):9–19.
- 61. Kikuchi T, Shibuya H, Jones Jxat. Molecular and biochemical characterization of an endo-beta-1, 3-glucanase from the pinewood nematode Bursaphelenchus xylophilus acquired by horizontal gene transfer from bacteria. Biochem J. 2005;389:117–25. pmid:15727561
- 62. Kikuchi T, Jones JT, Aikawa T, Kosaka H, Ogura N. A family of glycosyl hydrolase family 45 cellulases from the pine wood nematode Bursaphelenchus xylophilus. FEBS letters. 2004;572(1):201–5.
- 63. Mitreva-Dautova M, Roze E, Overmars H, de Graaff L, Schots A, Helder J, et al. A symbiont-independent endo-1, 4-β-xylanase from the plant-parasitic nematode Meloidogyne incognita. Mol Plant-Microbe In. 2006;19(5):521–9.
- 64. Adam MA, Phillips MS, Jones JT, Blok VC. Characterisation of the cellulose-binding protein Mj-cbp-1 of the root knot nematode, Meloidogyne javanica. Physiol Mol Plant P. 2008;72(1):21–8.
- 65. Eves-van den Akker S, Lilley CJ, Jones JT, Urwin PE. Identification and characterisation of a hyper-variable apoplastic effector gene family of the potato cyst nematodes. PLoS Pathog. 2014;10(9):e1004391. pmid:25255291
- 66. Gao B, Allen R, Maier T, Davis EL, Baum TJ, Hussey RS. Identification of putative parasitism genes expressed in the esophageal gland cells of the soybean cyst nematode Heterodera glycines. Mol Plant-Microbe In. 2001;14(10):1247–54.
- 67. Huang G, Gao B, Maier T, Allen R, Davis EL, Baum TJ, et al. A profile of putative parasitism genes expressed in the esophageal gland cells of the root-knot nematode Meloidogyne incognita. Mol Plant-Microbe In. 2003;16(5):376–81.
- 68. De Boer JM, Mcdermott JP, Wang X, Maier T, Qui F, Hussey RS, et al. The use of DNA microarrays for the developmental expression analysis of cDNAs from the oesophageal gland cell region of Heterodera glycines. Mol Plant Pathol. 2002;3(4):261–70. pmid:20569333
- 69. Noon JB, Hewezi TAF, Maier TR, Simmons C, Wei J-Z, Wu G, et al. Eighteen new candidate effectors of the phytonematode Heterodera glycines produced specifically in the secretory esophageal gland cells during parasitism. Phytopathology. 2015;(ja).
- 70. Dalzell JJ, McVeigh P, Warnock ND, Mitreva M, Bird DM, Abad P, et al. RNAi effector diversity in nematodes. PLoS Negl Trop Dis. 2011;5(6):e1176. pmid:21666793
- 71. Kumar M, Gantasala NP, T R, Thakur PK, Banakar P, Shukla RN, et al. De novo transcriptome sequencing and analysis of the cereal cyst nematode, Heterodera avenae. PloS one. 2014;9(5):e96311. pmid:24802510
- 72. Thorpe P, Mantelin S, Cock PJ, Blok VC, Coke MC, Eves-van den Akker S, et al. Genomic characterisation of the effector complement of the potato cyst nematode Globodera pallida. BMC genomics. 2014;15(1):923.
- 73. Danchin EGJ, Rosso M-N, Vieira P, de Almeida-Engler J, Coutinho PM, Henrissat B, et al. Multiple lateral gene transfers and duplications have promoted plant parasitism ability in nematodes. Proc Natl Acad Sci USA. 2010;107(41):17651–6. pmid:20876108
- 74. Bellafiore S, Shen Z, Rosso MN, Abad P, Shih P, Briggs SP. Direct identification of the Meloidogyne incognita secretome reveals proteins with host cell reprogramming potential. PLoS Pathog. 2008;4(10):e1000192. pmid:18974830
- 75. Wang J, Replogle AMY, Hussey R, Baum T, Wang X, Davis EL, et al. Identification of potential host plant mimics of CLAVATA3/ESR (CLE)-like peptides from the plant-parasitic nematode Heterodera schachtii. Mol Plant Pathol. 2011;12(2):177–86. pmid:21199567
- 76. Djamei A, Schipper K, Rabe F, Ghosh A, Vincon V, Kahnt J, et al. Metabolic priming by a secreted fungal effector. Nature. 2011;478(7369):395–8. pmid:21976020
- 77. Jones JT, Furlanetto C, Bakker E, Banks B, Blok V, Chen Q, et al. Characterization of a chorismate mutase from the potato cyst nematode Globodera pallida. Mol Plant Pathol. 2003;4(1):43–50. pmid:20569361
- 78. Lu S-W, Tian D, Borchardt-Wier HB, Wang X. Alternative splicing: A novel mechanism of regulation identified in the chorismate mutase gene of the potato cyst nematode Globodera rostochiensis. Mol Biochem Parasit. 2008;162(1):1–15.
- 79. Wang X, Allen R, Ding X, Goellner M, Maier T, de Boer JM, et al. Signal peptide-selection of cDNA cloned directly from the esophageal gland cells of the soybean cyst nematode Heterodera glycines. Mol Plant-Microbe In. 2001;14(4):536–44.
- 80. Félix M-A, Ashe A, Piffaretti J, Wu G, Nuez I, Bélicard T, et al. Natural and experimental infection of Caenorhabditis nematodes by novel viruses related to nodaviruses. PLoS Biol. 2011;9(1):e1000586. pmid:21283608
- 81. Wylie SJ, Luo H, Li H, Jones MGK. Multiple polyadenylated RNA viruses detected in pooled cultivated and wild plant samples. Arch Virol. 2012;157(2):271–84. pmid:22075920