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Genome Skimming: A Rapid Approach to Gaining Diverse Biological Insights into Multicellular Pathogens

  • Dee R. Denver ,

    denvedee@oregonstate.edu

    Affiliation Department of Integrative Biology, Oregon State University, Corvallis, Oregon, United States of America

  • Amanda M. V. Brown,

    Affiliation Department of Integrative Biology, Oregon State University, Corvallis, Oregon, United States of America

  • Dana K. Howe,

    Affiliation Department of Integrative Biology, Oregon State University, Corvallis, Oregon, United States of America

  • Amy B. Peetz,

    Affiliation USDA-ARS Horticultural Crops Research Laboratory, Corvallis, Oregon, United States of America

  • Inga A. Zasada

    Affiliation USDA-ARS Horticultural Crops Research Laboratory, Corvallis, Oregon, United States of America

Genome Skimming: A Rapid Approach to Gaining Diverse Biological Insights into Multicellular Pathogens

  • Dee R. Denver, 
  • Amanda M. V. Brown, 
  • Dana K. Howe, 
  • Amy B. Peetz, 
  • Inga A. Zasada
PLOS
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Introduction

Genomic data acquisition is now trivial for biologists. Yet, moving from millions of sequence reads to an assembled and annotated genome continues to pose a daunting challenge. The first animal genome sequenced arose from the free-living model nematode Caenorhabditis elegans [1]. This venture provided an unprecedented foundation for new insights into genome function and ‘omics tool development. However, the C. elegans endeavor has been tough to repeat, even with the advent of new high-throughput DNA sequencing technologies. For example, the first plant-parasitic nematode (PPN) genomes were published ten years after the C. elegans genome [2,3], and only five publication-quality PPN genomes are presently available [46].

Fig 1 overviews the course of a typical genome project. Millions of DNA sequences are initially collected in a matter of days, thanks to new DNA sequencing technologies. Early analytical phases (quality control and initial assembly) are also quick and usually straightforward. However, the subsequent computational stages (refining the assembly, gene prediction, and annotation) present significant bioinformatics bottlenecks. These lengthy in silico steps require multiple iterative stages of analysis, finally leading to a finished genome deemed “good enough” for publication. These latter stages often take years.

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Fig 1. Genome skimming schematic.

Boxes progressing diagonally from top left to bottom right show steps typical of conventional genome projects. Grey boxes show steps shared by genome skimming and conventional genome projects. Red boxes, arrows, and Xs show conventional genome project steps eliminated in the genome skimming approach. Green boxes show analyses specific to our genome skimming strategy.

https://doi.org/10.1371/journal.ppat.1005713.g001

The term “genome skimming” was recently coined [79] to describe shallow sequencing approaches aiming to uncover conserved ortholog sequences for phylogenomic studies. Here, we overview a genome skimming strategy applied to six PPN species but expand the scope beyond phylogenetics and toward diverse questions relating to pathogen function and biology. We demonstrate our strategy’s utility in rapidly revealing insights and new hypotheses relating to nematode genome structure, effector genes, and endosymbionts.

Genome Assembly Results

We applied our genome skimming strategy (Fig 1; see S1 Text) to six PPN species: Anguina agrostis, Globodera ellingtonae, Pratylenchus neglectus, P. penetrans, P. thornei, and Xiphinema americanum. Five of these species are in the “top ten” list of nematode plant pathogens [10]. Our approach begins like most genome projects by creating a single unrefined assembly for each PPN that provides a reference set of sequences for subsequent study. The lengthy downstream bioinformatics steps of typical genome projects, however, were simply not done. After completing single-pass assemblies, we examined the basic properties of the assembled contigs (Table 1). Assemblies yielded between ~10,000 and ~50,000 contigs per PPN, with average n-fold DNA sequence coverage values ranging from 7.7X to 30.4X. With an average coarse genome size estimate of 107.1 Mb and average GC content of 40.5%, these 6 PPN genome assembly patterns are consistent with known nematode genome size ranges [11,12]. We note that our smallest estimate (38.5 Mb) came from X. americanum, whose relative in the family Longidoridae, Longidorus kuiperi, also has a small genome size estimate of 56.5 Mb [13]. The N50 statistic, a common statistical measure for average length of a set of sequences (see S1 Text for more detail) was 8,863 bp on average for the six PPN species analyzed. Since nematode genes average ~2–3 kb in length [1,11,12], the contigs resulting from our single-pass assembly are sufficiently long to be useful database resources for BLAST [14].

Characterizing Genomic Variation

Early genome sequencing initiatives focused on model organisms such as C. elegans, in which sequenced DNA came from highly inbred lab populations. Modern pathogen genomics, however, often requires analysis of natural populations in which numerous factors can lead to deviations from the genomic uniformity of an inbred lab culture. For example, pathogens may display population-level genetic variation, within-individual heterozygosity, and other deviations (e.g., polyploidy or interspecies hybridization). These pose potential challenges but also opportunities for discovery. Interspecies hybridization and associated genome admixture is of increasing relevance to natural parasite populations [15]. Meloidogyne incognita, the world’s most devastating PPN species, evolved through between-species hybridization, as evidenced by recent phylogenomic analyses and the complex ploidy state of its nuclear genome [2,16]. The extent of hybridization among PPN species, however, remains unclear.

We developed a simple BLASTN-based method to quickly screen for evidence of genomic variation, using a list of 65 conserved single copy orthologs found in the genomes of C. elegans and G. rostochiensis (S1 Table) and our PPN genome assemblies. G. rostochiensis orthologs were used as queries against our G. ellingtonae contig database; single hits were found for all orthologs in the latter species, suggesting a high degree of genomic uniformity in the sample sequenced for this species. For the other 5 PPN species, however, more variable results were observed (Fig 2A). The median number of orthologs was equal to 1 for 2 species (A. agrostis, X. americanum), with small variances in copy number among the 65 genes (0.56 for A. agrostis, 1.25 for X. americanum). This small variation likely reflects some small genetic variation among the nematodes sequenced and/or the occurrence of lineage-specific duplicates for some of the orthologs. The median number of orthologs detected was 2 for all 3 Pratylenchus species. For the P. penetrans sample, it was known that nematodes from many field populations were combined in the sample used for the Illumina run, and thus, this genomic diversity is reflected in the high variance in ortholog copy number calculated for this species (4.46). The sequenced DNA samples for P. neglectus and P. thornei, however, each came from single nematode populations. The variances for these two species (0.67 and 0.95, respectively) were similar to those calculated for A. agrostis and X. americanum. The median value of two copies per ortholog for P. neglectus and P. thornei, combined with their low variance, suggests possible tetraploidy in these species. This hypothesis is supported by cytological evidence collected nearly 50 years ago [17] suggesting tetraploidy for P. neglectus and diploidy for P. penetrans.

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Fig 2. Box and blob plots.

(A) Box plots reporting results for numbers of homologs detected for 65 highly conserved orthologs in 5 PPN species analyzed. Results for G. ellingtonae are not included because this species was found to encode a single homolog for all 65 orthologs. (B) and (C) Blob plot results for X. americanum and P. penetrans, respectively. Colors indicate BLAST matches to different species of bacteria.

https://doi.org/10.1371/journal.ppat.1005713.g002

Finding Effector Genes

Discovery and functional characterization of effector genes, whose products directly engage in attacks on host defenses, is a central aim of any pathogen genome project. Protein sequences for 10 effectors, well characterized in other PPN species (S2 Table), were used as TBLASTN queries to screen our PPN contig databases for homologous matches. Our search revealed 42 matches (out of 60 possible) distributed across the PPN genomes (Table 1). As expected, more hits were observed in the 5 tylenchid PPN species analyzed (ranging from 6 to 8) compared to the very distantly related X. americanum, in which only 3 hits were observed. These 3 genes (annexin, β-1,4-endoglucanase, peroxiredoxin) were found in all 5 of the other species studied; a previous study revealed evidence for an expressed endoglucanase effector in X. index [18], a congener of X. americanum. The 3 X. americanum hit e-values (averaging 7.1 E-30) and hit lengths (averaging 459 bp) were larger and shorter, respectively, compared to averages for these 3 genes in the other 5 species (1.0 E-42, 632 bp). The addition of a simple single BLAST step to our genome skimming strategy quickly revealed the presence of numerous putative effector genes in the PPN species, though follow-up experimentation and analysis remains necessary to evaluate whether or not bona fide effectors are encoded by the DNA sequences identified.

Discovering Endosymbionts

Bacterial endosymbionts, such as Wolbachia spp., are well known and widespread components of diverse arthropods. Genome sequencing efforts in filarial nematode species revealed the presence of Wolbachia, which functions as an obligate mutualist in these pathogens of animals and humans [19,20].

We combined “Blob plot” approaches [21] with BLAST to uncover bacterial genomes associated with our PPN species. For the X. americanum analysis, evidence for its known endosymbiont Xiphinematobacter sp. [22] was observed as expected (Fig 2B). This genome-skimming result led to the hypothesis that the contigs in this blob constituted the Xiphinematobacter sp. genome. Follow-up bioinformatics, functional genomics, and fluorescence in situ hybridization (FISH) microscopy work supported this hypothesis and suggested that the endosymbiont functions as a nutritional mutualist with its nematode host [23].

A second interesting case was P. penetrans, in which 1,593 contigs matched bacterial DNA of diverse origins. Although many of these sequences contained high %GC, which were likely environmental contaminants (Fig 2C), two bacterial blobs of higher %AT were found containing contigs matching DNA of the known endosymbionts Wolbachia sp. and Cardinium sp. The only PPN previously reported to harbor Wolbachia is Radopholus similis [24]. A P. penetrans contig matched the 16S rDNA gene for Wolbachia in R. similis at 98% identity. Further bioinformatic and FISH work is underway to validate and build upon these initial endosymbiosis hypotheses arising from the P. penetrans genome skimming data.

Conclusions

Genome skimming provides a rapid and affordable avenue for biological inquiry and hypothesis generation that avoids the time delays that accompany most genomic endeavors. A single-pass assembly followed by BLAST-based and other simple analyses revealed evidence for potential genomic hybridization, effector genes, and endosymbionts in the PPN genomes studied. Although genome skimming provides an effective approach to hypothesis generation, follow-up work remains necessary for hypothesis evaluation. Genome skimming alone will not suffice for biological questions requiring gene prediction and annotation (e.g., patterns of gene family expansion, instances of horizontal gene transfer). Nonetheless, our genome skimming pilot experiment provided quick and exciting biological insights and community genomic resources, essentially doubling the number of PPN species for which published genome sequence resources are available. How might our understanding of nematode pathogens change if genome skimming were applied to 600 PPN species instead of 6?

Supporting Information

S1 Table. Conserved Orthologs Used in Genomic Variation Analysis.

https://doi.org/10.1371/journal.ppat.1005713.s002

(DOCX)

S2 Table. Effector Protein Sequences Used in BLAST Analysis.

https://doi.org/10.1371/journal.ppat.1005713.s003

(DOCX)

Acknowledgments

Thanks go to the Oregon State University Center for Genome Research and Biocomputing for Illumina DNA sequencing and bioinformatics support. We thank Wendy S. Phillips for helpful comments and bioinformatics contributions. Nematode samples were kindly provided by Dr. Steve Alderman and Dr. Guiping Yan.

References

  1. 1. C. elegans sequencing consortium. Genome sequence of the nematode C. elegans: a platform for investigating biology. The C. elegans Sequencing Consortium [published erratum appears in Science 1999 Jan 1;283(5398):35]. Science (80-). 1998;282: 2012–2018. pmid:9851916
  2. 2. 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: 909–915. pmid:18660804
  3. 3. 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 U S A. 2008;105: 14802–14807. pmid:18809916
  4. 4. Kikuchi T, Cotton JA, Dalzell JJ, Hasegawa K, Kanzaki N, McVeigh P, et al. Genomic insights into the origin of parasitism in the emerging plant pathogen bursaphelenchus xylophilus. PLoS Pathog. 2011;7(9):e1002219. pmid:21909270
  5. 5. Cotton JA, Lilley CJ, Jones LM, Kikuchi T, Reid AJ, 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: R43. pmid:24580726
  6. 6. Schaff JE, Windham E, Graham S, Crowell R, Scholl EH, Wright GM, et al. The plant parasite Pratylenchus coffeaecarries a minimal nematode genome. Nematology. Brill; 2015;17: 621–637.
  7. 7. Weitemier K, Straub SCK, Cronn RC, Fishbein M, Schmickl R, McDonnell A, et al. Hyb-Seq: Combining Target Enrichment and Genome Skimming for Plant Phylogenomics. Appl Plant Sci. 2014;2: 1400042.
  8. 8. Ripma LA, Simpson MG, Hasenstab-Lehman K. Geneious! Simplified genome skimming methods for phylogenetic systematic studies: A case study in Oreocarya (Boraginaceae). Appl Plant Sci. 2014;2: 1400062.
  9. 9. Malé PJG, Bardon L, Besnard G, Coissac E, Delsuc F, Engel J, et al. Genome skimming by shotgun sequencing helps resolve the phylogeny of a pantropical tree family. Mol Ecol Resour. 2014;14: 966–975. pmid:24606032
  10. 10. Jones JT, Haegeman A, Danchin EGJ, Gaur HS, Helder J, Jones MGK, et al. Top 10 plant-parasitic nematodes in molecular plant pathology. Mol Plant Pathol. 2013;14: 946–961. pmid:23809086
  11. 11. Coghlan A. Nematode genome evolution. WormBook. 2005; 1–15.
  12. 12. Bird DM, Williamson VM, Abad P, McCarter J, Danchin EGJ, Castagnone-Sereno P, et al. The genomes of root-knot nematodes. Annu Rev Phytopathol. 2009;47: 333–351. pmid:19400640
  13. 13. Leroy S, Bouamer S, Morand S, Fargette M. Genome size of plant-parasitic nematodes. Nematology. 2007;9: 449–450.
  14. 14. 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 Research. 1997. pp. 3389–3402. pmid:9254694
  15. 15. King KC, Stelkens RB, Webster JP, Smith DF, Brockhurst MA. Hybridization in Parasites: Consequences for Adaptive Evolution, Pathogenesis, and Public Health in a Changing World. PLoS Pathog. 2015;11: e1005098. pmid:26336070
  16. 16. Lunt DH, Kumar S, Koutsovoulos G, Blaxter ML. The complex hybrid origins of the root knot nematodes revealed through comparative genomics. PeerJ. 2014;2: e356. pmid:24860695
  17. 17. Roman J, Triantaphyllou AC. Gametogenesis and reproduction of seven species of pratylenchus. J Nematol. 1969;1: 357–62. http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=2617843&tool=pmcentrez&rendertype=abstract pmid:19325697
  18. 18. Furlanetto C, Cardle L, Brown DJF, Jones JT. Analysis of expressed sequence tags from the ectoparasitic nematode Xiphinema index. Nematology. 2005;7: 95–104.
  19. 19. Foster J, Ganatra M, Kamal I, Ware J, Makarova K, Ivanova N, et al. The Wolbachia genome of Brugia malayi: Endosymbiont evolution within a human pathogenic nematode. PLoS Biol. 2005;3: 0599–0614.
  20. 20. Fenn K, Blaxter M. Wolbachia genomes: Revealing the biology of parasitism and mutualism. Trends in Parasitology. 2006; 22(2):60–5. pmid:16406333
  21. 21. Kumar S, Jones M, Koutsovoulos G, Clarke M, Blaxter M. Blobology: exploring raw genome data for contaminants, symbionts and parasites using taxon-annotated GC-coverage plots. Front Genet. 2013;4: 237. pmid:24348509
  22. 22. Vandekerckhove TTM, Willems A, Gillis M, Coomans A. Occurrence of novel verrucomicrobial species, endosymbiotic and associated with parthenogenesis in Xiphinema americanum-group species (Nematoda, Longidoridae). Int J Syst Evol Microbiol. 2000;50: 2197–2205. pmid:11155997
  23. 23. Brown AM V, Howe DK, Wasala SK, Peetz AB, Zasada IA, Denver DR. Comparative Genomics of a Plant-Parasitic Nematode Endosymbiont Suggest a Role in Nutritional Symbiosis. Genome Biol Evol. 2015;7: 2727–46. pmid:26362082
  24. 24. Haegeman A, Vanholme B, Jacob J, Vandekerckhove TTM, Claeys M, Borgonie G, et al. An endosymbiotic bacterium in a plant-parasitic nematode: Member of a new Wolbachia supergroup. Int J Parasitol. 2009;39: 1045–1054. pmid:19504759