Improved 18S small subunit rDNA primers for problematic nematode amplification

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VOLUME 50 , ISSUE 4 (December 2018) > List of articles

Improved 18S small subunit rDNA primers for problematic nematode amplification

L. K. Carta * / S. Li

Keywords : Barcode, Molecular sequence, Nematode identification

Citation Information : Journal of Nematology. Volume 50, Issue 4, Pages 533-542, DOI: https://doi.org/10.21307/jofnem-2018-051

License : (PUBLISHER)

Published Online: 03-December-2018

ARTICLE

ABSTRACT

The 18S small subunit (SSU) ribosomal DNA sequence is one of the most useful molecular loci for identification and phylogeny reconstruction of agriculturally important nematodes. Various pairs of universal primers have been developed in the past to amplify short and long nematode sequences. However, certain nematode taxa were not readily amplified and/or sequenced with the existing primer tools. Frequently, the center region of a roughly 1,000 nucleotide segment would be lost. Therefore new primers were developed based on a very large 276 taxon alignment of 124 agriculturally important nematode species, and tested on problematic nematode taxa such as Aphelenchoides, Bursaphelenchus, Ditylenchus, and Panagrolaimus. New primers and protocols are provided for successful generation of sequences useful in future investigations of nematode systematics.

Graphical ABSTRACT

The 18S small subunit (SSU) ribosomal DNA (rDNA) sequence is one of the most useful molecular loci for identifying agricultural invertebrates (Kiewnick et al., 2016). Small segments have been successfully used as barcodes (Floyd et al., 2002; Powers, 2004), and longer segments are standard molecular sequences for deep level phylogenetics. Single or multiple pairs of primers were reported to amplify nematode 18S SSU rDNA from different taxon templates (Table 1). Although the amplifications by these primers have been reproducible in other studies, no single or multiple universal pairs of primers applicable across all nematode taxa have been reported. The challenges remain for what primers and strategies should be selected, particularly when the specimens are from unknown species with limited amounts of extracted DNA. Three pairs of universal 18S SSU rDNA primers that have been successfully used for many years to amplify a relatively long sequence with diverse nematodes (Thomas et al., 1997) (Table 1) for direct de novo sequencing of rDNA have been problematic for certain nematode taxa in our laboratory. The PCR failures with the primers were seen in Acrobeloides, Bursaphelenchus, and Ditylenchus and partially in Laimaphelenchus/Aphelenchoides heidelbergi (Carta et al., 2016) and other taxa. The failures were seen with the G18S4 and 18P primer pair (Table 1) as well (Blaxter et al., 1998). Sequencing failures were also observed in many cases even if all of the three primer pairs worked well enough to amplify a PCR product. Also, the use of multiple standard primer pairs consumed too much sample genomic DNA (2 µl of nematode lysate for each set) to satisfactorily amplify 18S rDNA when specimen DNA was in limited supply. Therefore, in this work we selected new primers from conserved DNA sequences in a multiple-sequence alignment of many plant parasitic and saprophytic nematodes, developed protocols, and successfully tested them on multiple, difficult-to-amplify specimens.

Table 1

Single pair or multiple pairs of primers frequently used for the amplification of near-full length 18S SSU rDNA.

10.21307_jofnem-2018-051-t001.jpg

Materials and methods

Primer design

Designing new consensus 18S primers was initially based on a multiple alignment of 266 18S sequences extracted from GenBank, representing 124 nematode species across the Nematoda, including plant parasitic and non-parasitic species, with some specifically related to address the issues above. Ultimately, 276 18S sequences from major plant parasitic nematodes, including Anguina, Aphelenchoides, Bursaphelenchus, Ditylenchus, Hemicycliophora, Heterodera, Longidorus, Meloidogyne, Mesocriconema, Nacobbus, Paratrichodorus, Pratylenchus, Rotylenchulus, Trichodorus, Tylenchulus, and Xiphinema, were further aligned with Clustal W (Thompson et al., 1994) in Geneious ver 8.1.7 (BioMatters, Auckland, New Zealand) to create consensus 18S primers. The strategy employed was to select primers, 19 to 36 nt long with a high melting temperature (Tm) around 60°C and without dimers and hairpins, as their presence can lead to poor or no yield of PCR product. Primers selected based on this strategy were listed in Table 2.

Table 2

Primers used for PCR and sequencing.

10.21307_jofnem-2018-051-t002.jpg

DNA analysis

Specimens were mechanically disrupted in 20 µl of extraction buffer (Thomas et al., 1997) then stored in PCR tubes at –80°C until needed. Extracts were prepared from thawed pools by incubating the tubes at 60°C for 60 min., followed by 95°C for 15 min. to deactivate proteinase K. Two microliters of the extract was used for each 25 µl PCR reaction within a Bio-Rad MJ Mini or C1000 Touch gradient thermal cycler (Bio-Rad Laboratories, Hercules, CA):

  1. TaKaRa Ex Taq

    • 10XEx Taq Buffer 5 μl

    • dNTP mixture (2.5 mM each) 4 μl

    • 10 μM forward primer 1 μl

    • 10 μM reverse primer 1 μl

    • TaKaRa Ex Taq (5 units/μl) 0.25 μl

    • 18S template DNA (>1,000bp) 4 μl

    • Sterilized distilled water up to 34.75 μl

  2. Invitrogen Platinum Taq

    • 10X PCR Buffer, – Mg 2.5 μl

    • 50 mM MgCl2 0.75 μl

    • 10 mM dNTP mix 0.5 μl

    • 10 μM forward prime 0.5 μl

    • 10 μM reverse primer 0.5 μl

    • 18S template DNA(<1,000bp) 2 μL

    • Platinum™ Taq 0.1 μl

    • Water, nuclease-free to 18.15 μl

  3. Phusion Taq

    • 5X Phusion HF Buffer 10 µl

    • 10 mM dNTPs 1 µl

    • 10 µM forward primer 2.5 µl

    • 10 µM reverse primer 2.5 µl

    • 18S template DNA 4 µl

    • Phusion DNA polymerase 0.5 µl

    • Water added up to 29.5 µl

One of the advantages of Phusion Taq is that its PCR is very rapid and can be done in less than 2 hr. It tends to generate multiple bands and the detergent in the buffer may interrupt sequencing downstream. Further evaluation may be advisable in particular circumstances.

PCR conditions

For TaKaRa Ex Taq and 18S Template DNA (>1,000bp): 95°C for 3 min, 5X (94°C for 30′, 45°C for 40′, 72°C for 2 min), 40X (94°C for 30′,Ta (°C) for 40′, 72°C for 2 min), 72°C for 5 min, 4°C until finish.

For Invitrogen Platinum™ Taq and 18S Template DNA (<1,000bp): 95°C for 3 min, 35X (94°C for 30′, Ta (°C) for 40′, 72°C for 70′), 72°C for 5 min, 4°C until finish

For Phusion and 18S Template DNA (>1,000bp): 98°C for 30′, 35X (98°C for 10′, 59°C for 30′, 72°C for 90′), 72°C for 2 min, 4°C until finish.

PCR products were visualized with the Lonza FlashGelTM DNA system (VWR International, Radnor, PA) and then treated with ExoSAP-IT reagent (Affymetrix, Inc, Santa Clara, CA) according to the manufacturer’s protocol. DNA sequencing was performed with an ABI BigDye Terminator v3.1 kit and in an ABI 3730xl DNA Analyzer (Applied Biosystems, Foster City, CA, USA) owned by the USDA Systematic Entomology Lab, Beltsville, MD.

Results and discussion

The G18S4 universal primer is a critical forward primer used for the 18SF-Cocktail (Thomas, 2011) paired with 18P (Table 1), and is comparable to the new 18S-CL-F3 and 18S-CL-F primers (Tables 1,2):

10.21307_jofnem-2018-051-ut001.jpg

To study which primer among these three is a better candidate, the reverse primer D2AR, with no primer-template mismatches in Aphelenchoides bicaudatus, Bursaphelenchus sp., Ditylenchus sp., and Panagrolaimus sp., was selected to minimize any discriminations in PCR amplification by a reverse primer, as shown in Figure 1. G18S4 amplified 18S and ITS rDNA of Ditylenchus sp., weakly amplified DNA of Panagrolaimus sp. and Aphelenchoides bicaudatus, but did not amplify Bursaphelenchus sp. DNA. The 18S-CL-F primer amplified the templates for Bursaphelenchus sp. but not amplify Ditylenchus sp. or Panagrolaimus sp., and weakly amplified the Aphelenchoides bicaudatus template. In contrast, the 18S-CL-F3 primer amplified the templates for these four taxa without exception.

Figure 1

Position diagram of 18S primers constructed with pDRAW32 DNA analysis by AcaClone software. http://www.acaclone.com/ Primer pairs in green and blue are older universal primers (Table 2), and those in red were generated in this study (Table 2).

10.21307_jofnem-2018-051-f001.jpg

After aligning the sequence results from Figure 1 with these three primer sequences above, it is evident that the lack of the 3′ end sequence (solid line above) in G18S4 (panel A) made this primer incapable of amplifying the 18S-ITS templates of Panagrolaimus sp. (lane 1), Aphelenchoides bicaudatus (lane 2) and Bursaphelenchus sp. (lane 3). On the other hand, removal of the 5′ end sequence (dash line above) in 18S-CL-F (panel B) made it incapable of amplifying the templates of Panagrolaimus sp. (lane 1), Aphelenchoides bicaudatus (lane 2), and Ditylenchus sp. (lane 4) (Fig. 2). While having more bases to prime than G18S4 and 18S-CL-F with these terminal sequences, the 18S-CL-F3 primer executed the amplifications very well. When PCR primers are designed, the sequence at the 3′ end generally commands more attention than at the 5′ end (Kwok et al.,1990; Onodera and Melcher, 2004), however, our results indicate the sequence at the 5′ end is equally critical. Taken together, these results suggest that 18S-CL-F3 is a better forward primer candidate than either G18S4 or 18S-CL-F to reliably generate a robust amplicon from this region of rDNA.

Figure 2

Comparison of 18S primers by the PCR amplification. (A) G18S4 and D2AR; (B) 18S-CL-F and D2AR; (C) 18S-CL-F3 and D2AR; M: DNA ladder (0.1-4.0kb); 1: Panagrolaimus sp. Idaho; 2: Aphelenchoides bicaudatus, Maryland; 3: Bursaphelenchus sp. MX; 4: Ditylenchus sp. Idaho; NC was the negative control. The PCRs were performed with TaKaRa Ex Taq as described in Materials and Methods and the conditions: 95°C for 5 min, 35X (95°C for 30′, 50°C for 45′, 72°C for 2 min 30′), 72°C for 2 min, 4°C until finish. An approximately 2,900 nt amplicon was generated.

10.21307_jofnem-2018-051-f002.jpg

Forward primer 18S-CL-F3 was not only employed successfully for these four taxa but also for many more taxa than for 18S-CL-F (Table 3). The results in Table 3 also show that 18S-CL-F3 tolerated a wider and higher temperature range (50-58°C) than 18S-CL-F. The 18S-CL-F3 forward primer paired with universal reverse primers successfully amplified not only short length (500-1,000bp) but also long (1,000-2,900bp) 18S templates (Table 3 and Fig. 2). In addition to the taxa in Table 3, Xiphinema, Hoplolaimus, Helicotylenchus, and Criconemoides were also tested successfully with the 18S-CL-F3 primer (data not shown).

Table 3

Taxa tested successfully with our newly designed ribosomal primers or paired with universal ribosomal primers.

10.21307_jofnem-2018-051-t003.jpg

Generally, one to three universal primer sets are needed for near-full length 18S sequence (Table 1) and one universal primer set for the ITS region (Vrain et al., 1992). However, by using 18S-CL-F3, only one primer set was sufficient to cover both 18S and ITS rDNA regions. Additionally a single primer set, 18S-CL-F2 or ITS-CL-F2, could be used with either 28S primers 1032R or 1006R (Table 2), respectively, to amplify rDNA that previously needed two sets for the ITS (Vrain et al., 1992) or the D1D2D3 regions of 28S (Nunn, 1992).

The 18S primers, Tyl2F and Tyl4R were designed to detect plant parasitic and fungivorous nematodes by PCR-Denaturing Gradient Gel Electrophoresis (PCR-DGGE) (Kushida, 2013). While using PCR-DGGE can reduce cost and time, it provides very limited sequence information. The forward Tyl2F primer is positioned approximately 345bp after the new 18S-CL-F3, and paired with Tyl4R it generates only a 450bp 18S fragment. This contrasts with the new 18S-CL-F3 primer, capable of amplifying near-full length 18S to ITS rDNA.

The degenerate primer Nem_SSU_F74 (Donn et al. (2011) was also designed to remedy 5′ primer mismatches for problematic and unknown taxa, but degenerate primers may still underperform for certain taxa. However, these new consensus primers have less bias and improved fidelity to reveal sequences that cover all nematodes across the Tylenchida and Rhabditida. The new 18S-CL-F3 primer binds to position 964-989 on the reference C. elegans sequence X03680, downstream of Nem_SSU_F74 that binds to positions 1007-1026 on the C. elegans reference sequence (Fig. 2). The new 18S-CL-F3 primer also lacks secondary structure which is often detrimental to PCR amplification, and its presence may compromise performance of the Nem_SSU_F74 primer. The lesser sensitivity of primer Nem_SSU_F74 also requires at least ten times more DNA (1-5 ng), than for the new 18S-CL-F3 (0.1 ng). Perhaps the greatest benefit of new primer 18S-CL-F3 is the capability of amplifying unusually long segments of rDNA, spanning from 18S into the entire ITS region when combined with other ribosomal primers, unlike the Nem_SSU_F74.

It should be noted that the utility of these ribosomal primers presented in this study are not limited to taxonomic identification and phylogenetic analysis using individual specimens. They can be used for biodiversity studies with metabarcoding from environmental DNA samples. The resulting amplicons can be sequenced using different NGS platforms, such as Illumina with short reads, and PacBio with long reads for long amplicons.

Acknowledgements

The authors thank Matt Lewis and Sonja Scheffer of the Insect Biocontrol Laboratory, USDA-ARS, Beltsville for use of their sequencing equipment and consultation regarding technical help. Mention of a trade name or commercial product in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the US Department of Agriculture. USDA is an equal opportunity provider and employer.

References


  1. Blaxter, M. L. , De Ley, P. , Garey, J. R. , Liu, L. X. , Scheldeman, P. , Vierstraete, A. , Vanfleteren, J. R. , Mackey, L. Y , Dorris, M. , Frisse, L. M. , Vida, J. T. , and Thomas, W. K. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392: 71–74.
  2. Carta, L. K. , Li, S. , Skantar, A. M. , and Newcombe, G. 2016. Morphological and molecular characterization of two Aphelenchoides endophytic in Poplar Leaves. Journal of Nematology 48: 28–33.
  3. Donn, S. , Neilson, R. , Griffiths, B. S. , and Daniell, T. J. 2011. Greater coverage of the phylum Nematoda in SSU rDNA studies. Biology and Fertility of Soils 47: 333–339.
  4. Floyd, R. , Abebe, E. , Papert, A. , and Blaxter, M. 2002. Molecular barcodes for soil nematode identification. Molecular Ecology 11: 839–850.
  5. Holterman, M. , van der Wurff, A. , van den Elsen, S. , van Megen, H. , Bongers, T. , Holovachov, O. , Bakker, J. , and Helder, J. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23: 1792–1800.
  6. Holterman, M. , Rybarczyk, K. , van Den Elsen, S. , van Megen, H. , Mooyman, P. , Peña Santiago, R. , Bongers, T. , Bakker, J. , and Helder, J. 2008. A ribosomal DNA-based framework for the detection and quantification of stress-sensitive nematode families in terrestrial habitats. Molecular Ecology Resources 8: 23–34.
  7. Joyce, S. A. , Reid, A. , Driver, F. , and Curran, J. 1994. Application of Polymerase Chain Reaction (PCR) methods to identification of entomopathogenic nematodes, in Burnell, A. M. , Ehlers, R. U. , and Masson, J. P. (Eds), COST 812 biotechnology: genetics of entomopathogenic nematode–bacterium complexes. proceedings of symposium and workshop, St. Patrick’s College, Maynooth, Co. Kildare, European Commission, DG XII, Ireland and Luxembourg, 178–187.
  8. Kanzaki, N. , and Futai, K. 2002. A PCR primer set for determination of phylogenetic relationships of Bursaphelenchus species within the xylophilus group. Nematology 4: 35–41.
  9. Kiewnick, S. , Buhlmann, A. , and Frey, J. E. 2016. DNA barcoding of invertebrate plant pests, in Boonham, N. , Tomlinson, J. , and Mumford, R. (Eds), Molecular methods in plant disease diagnostics, CABI, Wallingford, UK, 98–124.
  10. Kushida, A. 2013. Design and evaluation of PCR primers for denaturing gradient gel electrophoresis analysis of plant parasitic and fungivorous nematode communities. Microbes and Environments 28: 269–274.
  11. Kwok, S. , Kellogg, D. E. , McKinney, N. , Spasic, D. , Goda, L. , Levenson, C. , and Sninsky, J. J. 1990. Effects of primer-template mismatches on the polymerase chain reaction: human immunodeficiency virus type 1 model studies. Nucleic Acids Research 18: 999–1005.
  12. Lopez-Garcia, P. , Philippe, H. , Gail, F. , and Moreira, D. 2003. Autochthonous eukaryotic diversity in hydrothermal sediment and experimental microcolonizers at the Mid-Atlantic Ridge”, Proceedings of the National Academy of Sciences, Vol. 100: 697–702.
  13. Medlin, L. , Elwood, H. J. , Stickel, S. , and Sogin, M. L. 1988. The characterization of enzymatically amplified eukaryotic 16S-like rRNA-coding regions. Gene 71: 491–499.
  14. Mullin, P. G. , Harris, T. S. , and Powers, T. O. 2005. Phylogenetic relationships of Nygolaimina and Dorylaimina (Nematoda: Dorylaimida) inferred from small subunit ribosomal DNA sequences. Nematology 7: 59–79.
  15. Nunn, G. B. 1992. Nematode molecular evolution”, PhD dissertation, University of Nottingham, UK.
  16. Onodera, K. , and Melcher, U. 2004. Selection for 3′ end triplets for polymerase chain reaction primers. Molecular and Cellular Probes 18: 369–372.
  17. Olson, M. , Harris, T. , Higgins, R. , Mullin, P. , Powers, K. , Olson, S. , and Powers, T. O. 2017. Species delimitation and description of Mesocriconema nebraskense n. sp. (Nematoda: Criconematidae), a morphologically cryptic, parthenogenetic species from North American grasslands. Journal of Nematology 49: 42–66.
  18. Powers, T. 2004. Nematode molecular diagnostics: from bands to barcodes. Annual Review of Phytopathology 42: 367–383.
  19. Thomas, W. K. 2011. Molecular techniques., in International Seabed Authority (Eds), Marine benthic nematode molecular protocol handbook (nematode barcoding), Technical Study: No. 7, ISA Technical study series, 22–37 .
  20. Thomas, W. K. , Vida, J. T. , Frisse, L. M. , Mundo, M. , and Baldwin, J. G. 1997. DNA sequences from formalin-fixed nematodes: integrating molecular and morphological approaches to taxonomy. Journal of Nematology 29: 250–254.
  21. Thompson, J. D. , Higgins, D. G. , and Gibson, T. J. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22: 4673–4680.
  22. Vrain, T. C. , Wakarchuk, D. A. , Lévesque, A. C. , and Hamilton, R. I. 1992. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology 15: 563–573.
  23. Yaghoubi, A. , Pourjam, E. , Álvarez-Ortega, S. , Liébanas, G. , Atighi, M. R. , and Pedram, M. 2016. Discopersicus n. gen., a new member of the family Tylenchidae Örley, 1880 with detailed SEM study on two known species of the genus Discotylenchus Siddiqi, 1980 (Nematoda; Tylenchidae) from Iran. Journal of Nematology 48: 214–221.
  24. Zeng, Y. , Ye, W. , Tredway, L. , Martin, S. , and Martin, M. 2012. Description of Hemicaloosia graminis n. sp. (Nematoda: Caloosiidae) associated with turfgrasses in North and South Carolina, USA. Journal of Nematology 44: 134–141.
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FIGURES & TABLES

Figure 1

Position diagram of 18S primers constructed with pDRAW32 DNA analysis by AcaClone software. http://www.acaclone.com/ Primer pairs in green and blue are older universal primers (Table 2), and those in red were generated in this study (Table 2).

Full Size   |   Slide (.pptx)

Figure 2

Comparison of 18S primers by the PCR amplification. (A) G18S4 and D2AR; (B) 18S-CL-F and D2AR; (C) 18S-CL-F3 and D2AR; M: DNA ladder (0.1-4.0kb); 1: Panagrolaimus sp. Idaho; 2: Aphelenchoides bicaudatus, Maryland; 3: Bursaphelenchus sp. MX; 4: Ditylenchus sp. Idaho; NC was the negative control. The PCRs were performed with TaKaRa Ex Taq as described in Materials and Methods and the conditions: 95°C for 5 min, 35X (95°C for 30′, 50°C for 45′, 72°C for 2 min 30′), 72°C for 2 min, 4°C until finish. An approximately 2,900 nt amplicon was generated.

Full Size   |   Slide (.pptx)

REFERENCES

  1. Blaxter, M. L. , De Ley, P. , Garey, J. R. , Liu, L. X. , Scheldeman, P. , Vierstraete, A. , Vanfleteren, J. R. , Mackey, L. Y , Dorris, M. , Frisse, L. M. , Vida, J. T. , and Thomas, W. K. 1998. A molecular evolutionary framework for the phylum Nematoda. Nature 392: 71–74.
  2. Carta, L. K. , Li, S. , Skantar, A. M. , and Newcombe, G. 2016. Morphological and molecular characterization of two Aphelenchoides endophytic in Poplar Leaves. Journal of Nematology 48: 28–33.
  3. Donn, S. , Neilson, R. , Griffiths, B. S. , and Daniell, T. J. 2011. Greater coverage of the phylum Nematoda in SSU rDNA studies. Biology and Fertility of Soils 47: 333–339.
  4. Floyd, R. , Abebe, E. , Papert, A. , and Blaxter, M. 2002. Molecular barcodes for soil nematode identification. Molecular Ecology 11: 839–850.
  5. Holterman, M. , van der Wurff, A. , van den Elsen, S. , van Megen, H. , Bongers, T. , Holovachov, O. , Bakker, J. , and Helder, J. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23: 1792–1800.
  6. Holterman, M. , Rybarczyk, K. , van Den Elsen, S. , van Megen, H. , Mooyman, P. , Peña Santiago, R. , Bongers, T. , Bakker, J. , and Helder, J. 2008. A ribosomal DNA-based framework for the detection and quantification of stress-sensitive nematode families in terrestrial habitats. Molecular Ecology Resources 8: 23–34.
  7. Joyce, S. A. , Reid, A. , Driver, F. , and Curran, J. 1994. Application of Polymerase Chain Reaction (PCR) methods to identification of entomopathogenic nematodes, in Burnell, A. M. , Ehlers, R. U. , and Masson, J. P. (Eds), COST 812 biotechnology: genetics of entomopathogenic nematode–bacterium complexes. proceedings of symposium and workshop, St. Patrick’s College, Maynooth, Co. Kildare, European Commission, DG XII, Ireland and Luxembourg, 178–187.
  8. Kanzaki, N. , and Futai, K. 2002. A PCR primer set for determination of phylogenetic relationships of Bursaphelenchus species within the xylophilus group. Nematology 4: 35–41.
  9. Kiewnick, S. , Buhlmann, A. , and Frey, J. E. 2016. DNA barcoding of invertebrate plant pests, in Boonham, N. , Tomlinson, J. , and Mumford, R. (Eds), Molecular methods in plant disease diagnostics, CABI, Wallingford, UK, 98–124.
  10. Kushida, A. 2013. Design and evaluation of PCR primers for denaturing gradient gel electrophoresis analysis of plant parasitic and fungivorous nematode communities. Microbes and Environments 28: 269–274.
  11. Kwok, S. , Kellogg, D. E. , McKinney, N. , Spasic, D. , Goda, L. , Levenson, C. , and Sninsky, J. J. 1990. Effects of primer-template mismatches on the polymerase chain reaction: human immunodeficiency virus type 1 model studies. Nucleic Acids Research 18: 999–1005.
  12. Lopez-Garcia, P. , Philippe, H. , Gail, F. , and Moreira, D. 2003. Autochthonous eukaryotic diversity in hydrothermal sediment and experimental microcolonizers at the Mid-Atlantic Ridge”, Proceedings of the National Academy of Sciences, Vol. 100: 697–702.
  13. Medlin, L. , Elwood, H. J. , Stickel, S. , and Sogin, M. L. 1988. The characterization of enzymatically amplified eukaryotic 16S-like rRNA-coding regions. Gene 71: 491–499.
  14. Mullin, P. G. , Harris, T. S. , and Powers, T. O. 2005. Phylogenetic relationships of Nygolaimina and Dorylaimina (Nematoda: Dorylaimida) inferred from small subunit ribosomal DNA sequences. Nematology 7: 59–79.
  15. Nunn, G. B. 1992. Nematode molecular evolution”, PhD dissertation, University of Nottingham, UK.
  16. Onodera, K. , and Melcher, U. 2004. Selection for 3′ end triplets for polymerase chain reaction primers. Molecular and Cellular Probes 18: 369–372.
  17. Olson, M. , Harris, T. , Higgins, R. , Mullin, P. , Powers, K. , Olson, S. , and Powers, T. O. 2017. Species delimitation and description of Mesocriconema nebraskense n. sp. (Nematoda: Criconematidae), a morphologically cryptic, parthenogenetic species from North American grasslands. Journal of Nematology 49: 42–66.
  18. Powers, T. 2004. Nematode molecular diagnostics: from bands to barcodes. Annual Review of Phytopathology 42: 367–383.
  19. Thomas, W. K. 2011. Molecular techniques., in International Seabed Authority (Eds), Marine benthic nematode molecular protocol handbook (nematode barcoding), Technical Study: No. 7, ISA Technical study series, 22–37 .
  20. Thomas, W. K. , Vida, J. T. , Frisse, L. M. , Mundo, M. , and Baldwin, J. G. 1997. DNA sequences from formalin-fixed nematodes: integrating molecular and morphological approaches to taxonomy. Journal of Nematology 29: 250–254.
  21. Thompson, J. D. , Higgins, D. G. , and Gibson, T. J. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Research 22: 4673–4680.
  22. Vrain, T. C. , Wakarchuk, D. A. , Lévesque, A. C. , and Hamilton, R. I. 1992. Intraspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology 15: 563–573.
  23. Yaghoubi, A. , Pourjam, E. , Álvarez-Ortega, S. , Liébanas, G. , Atighi, M. R. , and Pedram, M. 2016. Discopersicus n. gen., a new member of the family Tylenchidae Örley, 1880 with detailed SEM study on two known species of the genus Discotylenchus Siddiqi, 1980 (Nematoda; Tylenchidae) from Iran. Journal of Nematology 48: 214–221.
  24. Zeng, Y. , Ye, W. , Tredway, L. , Martin, S. , and Martin, M. 2012. Description of Hemicaloosia graminis n. sp. (Nematoda: Caloosiidae) associated with turfgrasses in North and South Carolina, USA. Journal of Nematology 44: 134–141.

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