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Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Specific scope

This standard describes a diagnostic protocol for Globodera rostochiensis and Globodera pallida1.

Specific approval and amendment

This Standard was developed under the EU DIAGPRO Project (SMT 4-CT98-2252) by partnership of contractor laboratories and intercomparison laboratories in European countries. Approved as an EPPO Standard in 2003–09. Revision approved in 2009–09.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Globodera rostochiensis and Globodera pallida (potato cyst nematodes) cause major losses in Solanum tuberosum (potato) crops (van Riel & Mulder, 1998). The infective second stage juveniles only move a maximum of about 1 m in the soil. Most movement to new localities is by passive transport. The main routes of spread are infested seed potatoes and movement of soil (e.g. on farm machinery) from infested land to other areas. Infestation occurs when the second-stage juvenile hatches from the egg and enters the root near the growing tip by puncturing the epidermal cell walls, and then internal cell walls, with its stylet. Eventually it begins feeding on cells in the pericycle, cortex or endodermis. The nematode induces an enlargement of root cells and breakdown of their walls to form a large, syncytial transfer cell. This syncytium provides nutrients for the nematode. Infested potato plants have a reduced root system and, because of the decreased water uptake, plant death can eventually occur.

Identity

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Name:Globodera rostochiensis (Wollenweber, 1923), Skarbilovich, 1959

Synonyms:Heterodera rostochiensis, Wollenweber, 1923; Heterodera schachtii solani Zimmerman, 1927; Heterodera schachtii rostochiensis (Wollenweber) Kemner, 1929

Taxonomic position: Nematoda, Tylenchida2, Heteroderidae

EPPO computer code: HETDRO

Phytosanitary categorization: EPPO A2 list No. 125, EU Annex designation I/A2.

Name: Globodera pallida (Stone, 1973)

Synonyms:Heterodera pallida Stone, 1973

Taxonomic position: Nematoda, Tylenchida2, Heteroderidae

EPPO code: HETDPA

Phytosanitary categorization: EPPO A2 list No. 124, EU Annex designation I/A2.

Detection

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Symptoms

Above-ground symptoms due to potato cyst nematodes are not specific and often go undetected. General symptoms include patches of poor growth in the crop, with plants sometimes showing yellowing, wilting or death of foliage; tuber size is reduced and roots are extensively branched with soil stuck to them. However, many other causes can lead to these symptoms. Plants should therefore be lifted for a visual check on the presence of cysts and young females on the roots, or a soil sample should be taken for testing. Young females and cysts are just visible to the naked eye as tiny white, yellow or brown eye pin-heads on the root surface (Figs 1 and 2). Detection by lifting plants is only possible for a short time as females mature into cysts and then can easily be lost at lifting, and it is time-consuming. Soil testing is therefore the best way of determining the presence of potato cyst nematodes.

image

Figure 1.  Potato roots infected by G. rostochiensis (Courtesy: Plant Protection Service, NL).

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Figure 2.  Broken cyst with eggs of G. pallida (Courtesy: Plant Protection Service, NL).

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Statutory sampling procedures

Details on sampling can be found in EPPO Standard PM 3/30 (OEPP/EPPO, 1991, under revision) or in the Council Directive 2007/33/EC of 11 June 2007 on control of potato cyst nematode (EU, 2007).

Extraction procedures

There are various processes for extracting cysts from the soil. Simple methods based on flotation can be as good as elutriation. For each method, a description is given in Appendix 1. Globodera cysts are generally round which distinguishes them from most other types of nematode cysts.

When moist soil samples are not immediately processed and viability tests are envisaged, they should be stored below 5°C as juveniles may hatch above 5°C. In addition soil samples should not be dried at a higher temperature than 25°C (and not lower than 40% air humidity) when viability tests are envisaged.

Bioassay

Another procedure to detect the nematodes is the bioassay (Appendix 2 test A).

Identification

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Second-stage juveniles and cysts should be obtained from soil, plant roots or tubers. The female colour at the appropriate stage can be used as an indication: a female which changes during maturation from white to yellow, then into a brown cyst, is G. rostochiensis, while one which changes from white directly to brown is G. pallida. Identification of cysts and other stages is in general based on a combination of morphological and morphometric characters and biochemical techniques. For light microscope identification, it is recommended to examine specimens mounted in fixative on microscope slides. A flow diagram indicating combinations of methods is given in Fig. 3.

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Figure 3.  Flow-diagram for identification of Globodera rostochiensis and G. pallida.

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Identification on the basis of morphological features

Identification of cyst and juveniles to genus level

To identify the cysts to genus level the key of Brzeski (1998), Baldwin & Mundo-Ocampo (1991), Wouts & Baldwin (1998) or Siddiqi (2000) should be used based on form of the cysts and the characteristics of the vulva-anus region.

Globodera cysts should meet the following characteristics. Cysts of Globodera are smoothly rounded with small projecting neck, no terminal cone present, diameter ± 450 μm, and have a tanned brown skin. Cuticle surface with zigzag pattern of ridges, a distinct D-layer is present. The perineal area consists of a single circumfenestration around the vulval slit, perineal tubercles on crescents near vulva. Anus subterminal without fenestra, vulva in a vulval basin, underbridge and bullae rarely present (Fig. 4). Eggs retained in cyst, no egg-mass present.

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Figure 4.  The perineal region of a Globodera cyst (after Fleming & Powers, 1998).

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Globodera juveniles should meet the following characteristics. The mobile second-stage juveniles of Globodera are vermiform, annulated and tapering at both ends. Within the genus Globodera, body length ranging from 445 to 510 μm, stylet length 19–25 μm, tail length 37–55 μm and a hyaline tail part of 21–31 μm.

Juvenile cyst nematodes may be found in soil with which extraction was performed for the detection of free-living nematodes. They can be distinguished from root-knot nematode juveniles (Meloidogyne spp.) by a more heavily sclerotised lip region, relatively large stylet, shape of the tail and more robust appearance (Fig. 5). In such cases, it is advised to perform an additional cyst extraction and to proceed with this diagnostic protocol.

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Figure 5.  difference between Meloidogyne and Globodera juveniles. Comparison between Meloidogyne fallax and G. pallida.

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Identification to species level

The identification of Globodera to species based on morphology can be difficult because of the observed variability of key characteristics. Therefore the use of a combination of cyst and second-stage juvenile characteristics is recommended for reliable identification.

G. rostochiensis and G. pallida are morphologically and morphometrically closely related (Stone, 1973a,b). Figure 6 presents some drawings of different stages of G. rostochiensis (Fig. 6A) and G. pallida (Fig. 6B). For cysts, the most important diagnostic differences are in the perineal area, i.e. number of cuticular ridges between vulva-anus and Granek’s ratio (Fig. 7A,B). The second-stage juvenile characteristics are stylet length and stylet knob shape (Table 1; Fig. 7C). As the range of values for each of these characteristics can overlap between species, care is needed. In such cases, confirmation with molecular techniques is recommended.

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Figure 6.  (A) G. rostochiensis. (A) Entire juvenile. (B) Head region of 2nd-stage juvenile. (C) 2nd-stage juvenile lateral field, mid-body. (D) Pharyngeal region of 2nd-stage juvenile. (E) Pharyngeal region of male. (F) Tail of male. (G) Lateral field of male, mid-body. (H) Entire cysts. (I) Head and neck of female. (J) Entire male. (After: C.I.H. Descriptions of Plant-Parasitic Nematodes, Set 2, No. 16). (B) G. pallida n. sp. Larva. (A) Entire. (B) Anterior. (C) Head. (D) Tail. (E) Lateral field mid-body region. (F) Lateral field tail. (G) Head en face at level of lips. (H) Head en face at level of base (after Stone, 1972).

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Figure 7.  (A) Perineal measurements for Globodera identification. (B) Vulval-anal ridge patterns for four Globodera species. (C) Stylets from four species of Globodera (after Flemming and Powers, 1998).

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Table 1.   Range and mean values of some essential characters of Globodera rostochiensis, G. pallida, G. tabacum (tabacum), G. achilleae and G. artemisiae as given in Baldwin & Mundo-Ocampo (1991), Brzeski (1998); Manduric & Anderson (2004) and Dobosz et al. (2006)
SpeciesJ2 Body length μmJ2 styletCyst measurements
Stylet knob width (μm)Shape of stylet knobs Length (μm)Number of cuticular ridges* between anus and vulval basin Granek’s ratio
  1. *In some cases the cuticular ridges may not be visible.

G. rostochiensis468 (425–505)3.2–4.0Anteriorly flattened to rounded, without forward projections19–23 (21.8)16–31 (>14)1.3–9.5 (>3)
G. pallida484 (440–525)4–5Forward projections22–24 (23.8)8–20 (<14)1.2–3.5 (<3)
G. tabacum (tabacum)477 (410–527)4–5Rounded23–2410–141–4.2 (<2.8)
G. achilleae492 (472–515)4–5Rounded to anchor shape24–26 (25)4–11 (<10)1.3–1.9 (mean 1.6)
G. artemisiae413 (357–490)3–5Rounded, anteriorly flattened, sometimes slightly indented18–29 (23)5 –16 0.8–1.7 (mean 1.0)

The three other Globodera species which could cause confusion during identification of potato cyst nematodes in Europe are G. achilleae (Golden & Klindic, 1973) Behrens, 19753, G. artemisiae (Eroshenko & Kazachenko, 1972) Behrens, 1975, and G. tabacum sensu lato. These first two species are not parasitic on potato, but recorded on Achillea millefolium and Artemisia vulgaris, respectively, in comparable agricultural areas. The G. tabacum species complex [G. tabacum tabacum (Lownsbery & Lownsbery, 1954) Skarbilovich, 1959; G. tabacum solanacearum (Miller & Gray, 1972) Behrens, 1975, and G. tabacum virginiae (Miller & Gray, 1972) Behrens, 1975] is found in North and Central America. In Southern Europe, G. tabacum tabacum is also present. It parasitizes Nicotiana tabacum (tobacco) and some other solanaceous plants (but not potato). See Table 1 and Fig. 7 for a morphometric and morphological comparison between potato cyst nematodes, G. achilleae, G. artemisiae and G. tabacum. See also Baldwin & Mundo-Ocampo (1991), Brzeski (1998) and Wouts & Baldwin, 1998) for more detailed information on other members of the Heteroderinae and identification keys.

Biochemical and molecular methods

As G. rostochiensis and G. pallida are morphologically closely related, many different biochemical techniques have been developed to distinguish the two species. Schots et al. (1992) were able to differentiate and quantify them using a set of three monoclonal antibodies. There were, however, cross reactivity problems between the antibodies for the two species.

Several scientists have developed PCR-based tests to separate the two potato cyst nematode species (e.g. Mulholland et al., 1996; Shields et al., 1996; Thiéry & Mugniéry,1996; Bulman & Marshall, 1997; Fullaondo et al., 1999; Fleming et al., 2000; Zouhar et al., 2000; Bates et al., 2002). Recommended tests are described in Appendix 3.

Isoelectric focusing (IEF) has proved to be sensitive enough to identify samples of potato cyst nematodes. During IEF, proteins are separated in a pH gradient and focused at the position in the gradient (the isoelectric point) where they become electrically neutral. Four species-specific proteins were located, pI 5.9 and 8.7 and pI 5.7 and 6.9. These proteins can be used to identify G. rostochiensis and G. pallida (Fleming & Marks, 1982; Karssen et al., 1995). Other techniques, such as RFLP analysis of total DNA (Burrows & Boffey, 1986), diagnostic probes (Marshall & Crawford, 1987; Burrows & Perry, 1988), and dot-blotting with specific probes (Marshall, 1993) have also proved to be useful to distinguish the two species. For a complete treatise on the use of immunology, protein electrophoresis, IEF and DNA, see Fleming et al. (2000). A comparison of IEF, ELISA and PCR used for the identification of potato cyst nematodes in field samples was made by Ibrahim et al. (2001).

Most techniques have, however, been developed especially to distinguish G. rostochiensis from G. pallida but have not been tested so far against species such as G. achilleae, G. tabacum or G. ‘mexicana’. This limitation should be noted. There may also be differences between European and non-European populations of the two species.

Identification of G. rostochiensis and G. pallida should preferably combine morphological and molecular methods (see Appendix 3 for molecular test). The tests used should be validated for specificity and cross reactions with related species.

Pathotypes

The term ‘pathotype’ is used by the International PCN Pathotype Scheme proposed by Kort et al. (1977) but is now considered too general. Some PCN populations cannot conclusively be assigned to the pathotypes described in this scheme. There are differences in virulence within the two PCN species, in particular between populations of G. pallida and they are of utmost importance in populations from South America, but identification at this level is not possible yet. Any population showing signs of a new virulence should be tested as soon as possible. In practice, the virulence of populations can be tested on a set of cultivars used in each country.

Testing the viability of eggs and juveniles

Testing of the viability of the eggs and juveniles may be required for regulatory purposes. This can be done by different methods.

  • 1
    Visual morphological determination of viability, a table with descriptions and figures is provided (Appendix 4). These observations require trained personnel.
  • 2
    Determination of viability with a bioassay. Two tests are described in Appendix 2. Such tests require more time to perform than visual morphological determination of viability and generally more time than determination of viability by hatching tests.
  • 3
    Determination of viability by hatching tests. Three tests are described in Appendix 5. Such tests require more time to perform than visual morphological determination of viability. When determining the viability with a hatching test, it should be noted that cysts which have formed recently may be dormant (e.g. when sampling is performed in autumn after potato harvest). In such situations cysts should be exposed to +4°C for at least 2–3 weeks.

Morphological determination of viability by staining with Meldola’s Blue is also possible but the chemical is not easily available so this technique is not described in the protocol.

Molecular tests are under development for testing the viability of eggs and juveniles but they are not available for routine testing at the moment.

Reference material

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Reference material can be obtained from:

Plant Protection Service, PO Box 9102, 6700 HC Wageningen (NL).

Food and Environmental Research Agency (FERA), Sand Hutton, York YO41 1LZ (GB).

Julius Kühn-Institut (JKI), Federal Research Centre for Cultivated Plants, Messeweg 11–12, 38104 Braunschweig (DE).

French National Institute for Agricultural Research (INRA) Biology of Organisms and Populations for Plant Protection Domaine de la Motte, BP 35327 35653 Le Rheu Cedex (FR).

Department of Plant Protection Biology – Nematology, Swedish University of Agricultural Sciences, Alnarp (SE).

Reporting and documentation

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Guidance on reporting and documentation is given in EPPO Standard PM7/77 (1) Documentation and reporting on a diagnosis.

Further information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Further information on this organism can be obtained from: L den Nijs and G Karssen, Plant Protection Service, National Reference Laboratory, PO Box 9102, 6700 HC Wageningen (NL); e-mail: l.j.m.f.den.nijs@minlnv.nl.

Footnotes
  • 1

    Use of brand names of chemicals or equipment in these EPPO Standards implies no approval of them to the exclusion of others that may also be suitable.

  • 2

    Recent development combining a classification based on morphological data and molecular analysis refer to ‘Tylenchomorpha’ (De Ley & Blaxter, 2004).

  • 3

    Krall (1978) considered G. millefolii (Kirjanova & Krall, 1965) Behrens, 1975 as species inquirenda, as the description was based on a single female. Brzeski (1998) reported on G. achilleae: ‘it may be conspecific with G. millefolii’. Additional research is needed to prove if G. achilleae is a junior synonym of G. millefolii.

  • 4

    They can be obtained from Ritter GmbH, Schwabenstraße 50–54, D-86836 Untermeitingen.

  • 5

    Based on ring testing between laboratories in The Netherlands, L. den Nijs.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

This protocol was originally drafted by: den Nijs L & Karssen G, Plant Protection Service, National Reference Laboratory, PO Box 9102, 6700 HC Wageningen, The Netherlands. It was revised by Anthoine G (French National Laboratory, Nematology section), Kox L and den Nijs L (Plant Protection Service, National Reference Laboratory, Wageningen (NL).

Methods performed in Germany and Austria provided by Kaemmerer D, Bayerische Landesanstalt für Landwirtschaft. Methods performed in Norway provided by Magnusson C. Methods performed in Sweden provided by Manduric S.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices
  • Baldwin JG & Mundo-Ocampo M (1991) Heteroderinae, cyst- and non-cyst-forming nematodes. In: Manual of Agricultural Nematology (Ed. NickleWR), pp. 275362. Marcel Dekker, New York, NY (US).
  • Bates JA, Taylor EJA, Gans PT & Thomas JE (2002) Determination of relative proportions of Globodera species in mixed populations of potato cyst nematodes using PCR product melting peak analysis. Molecular Plant Pathology 3, 153161.
  • Brzeski MW (1998) Nematodes of Tylenchina in Poland and Temperate Europe. Muzeum i Instytut Zoologii PAN, Warsaw (PL).
  • Bulman SR & Marshall JW (1997) Differentiation of Australasian potato cyst nematode (PCN) populations using the polymerase chain reaction (PCR). New Zealand Journal of Crop and Horticultural Science 25, 123129.
  • Burrows PR & Boffey SA (1986) A technique for the extraction and restriction endonuclease digestion of total DNA from Globodera rostochiensis and Globodera pallida second-stage juveniles. Revue de Nématologie 9, 199200.
  • Burrows PR & Perry RN (1988) Two cloned DNA fragments which differentiate Globodera pallida from G. rostochiensis. Revue de Nématologie 11, 441445.
  • De Ley P & Blaxter ML (2004) A new system for Nematoda: combining morphological characters with molecular trees, and translating clades into ranks and taxa. In: Proceedings of the Fourth International Congress of Nematology, 8–13 June 2002 Tenerife (ES) (Ed. CookR & HuntDJ), 865 pp. Brill, Leiden (NL).
  • Dobosz R, Obrepalska-Steplowska A & Kornobis S (2006) Globodera artemisiae (Eroshenko et Kazachenko, 1972) (Nematoda: Heteroderidae) from Poland. Journal of Plant Protection Research 46, 403407.
  • EU (2007) Council Directive 2007/33/EC of 11 June 2007 on control of potato cyst nematode and repealing Directive 69/465/EEC. Official Journal of the European Communities L156, 1222.
  • Fleming CC & Marks RJ (1982) A method for quantitative estimation of Globodera rostochiensis and Globodera pallida in mixed species samples. Record of Agricultural Research of the Department of Agriculture for Northern Ireland 30, 6770.
  • Fleming CC & Powers TO (1998) Potato cyst nematode diagnostics: morphology, differential hosts and biochemical techniques. In: Potato Cyst Nematodes, Biology, Distribution and Control (Ed. MarksRJ & BrodieBB), pp. 91114. CAB International, Wallingford (GB).
  • Fleming CC & Powers TO (1998) Potato cyst nematodes, species, pathotypes and virulence concepts. In: Potato cyst nematodes: biology, distribution and control (Eds MarksRJ & BrodieBB), pp. 51115. CAB International, Wallingford (GB).
  • Fleming CC, Rao J, Moreland B, Craig D & Turner SJ (2000) Diagnostics of cyst nematodes and tephritid fruit flies using mitochondrial and ribosomal DNA. Bulletin OEPP/EPPPO Bulletin 30, 585590.
  • Folkertsma RT, Van Der Voort JNAMR, Van Gent-Pelzer MPE, Den Groot KE, Van Den Bos WJR, Schots A, Bakker J & Gommers FJ (1994) Inter- and intraspecific variation between populations of Globodera rostochiensis and G. pallida revealed by random amplified polymorphic DNA. Phytopathology 84, 807811.
  • Fullaondo A, Barrena E, Viribay M, Barrena I, Salazar A & Ritter E (1999) Identification of potato cyst nematode species Globodera rostochiensis and G. pallida by PCR using specific primer combinations. Nematology 1, 157163.
  • Golden AM & Klindic O (1973) Heterodera achilleae n. sp. (Nematoda: Heteroderidae) from yarrow in Yugoslavia. Journal of Nematology 5, 196201.
  • Holterman M, Van Der Wurff A, Van Den Elsen S, Van Megen H, Holovachov O, Bakker J & Helder J (2006) Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution towards crown clades. Molecular Biology and Evolution 23, 17921800.
  • Ibrahim SK, Minnis ST, Barker ADP, Russell MD, Haydock PPJ, Evans K, Grove IG, Woods SR & Wilcox A (2001) Evaluation of PCR, IEF and ELISA techniques for the detection and identification of potato cyst nematodes from field soil samples in England and Wales. Pest Management Science 57, 10681074.
  • Karssen G, Van Hoenselaar T, Verkerk-Bakker B & Janssen R (1995) Species identification of cyst and root-knot nematodes from potato by electrophoresis of individual females. Electrophoresis 16, 105109.
  • Kort J, Ross H, Rumpenhorst HJ & Stone AR (1977) An international scheme for identifying and classifying pathotypes of potato cyst-nematodes Globodera rostochiensis and G. pallida. Nematologica 23, 333339.
  • Krall E (1978) Compendium of cyst nematodes in the USSR. Nematologica 23, 311332.
  • Manduric S & Anderson S (2004) The identification of Swedish Globodera (Nematoda, Heteroderidae) populations, following comparisons with known populations of G. artemisiae (Eroshenko and Kazachenko, 1972) Behrens, 1975. Russian Journal of Nematology 12, 3944.
  • Marshall JW (1993) Detecting the presence and distribution of Globodera rostochiensis and G. pallida mixed populations in New Zealand using DNA probes. New Zealand Journal of Crop and Horticultural Science 21, 219223.
  • Marshall JW & Crawford AM (1987) A cloned DNA fragment that can be used as a sensitive probe to distinguish Globodera pallida from Globodera rostochiensis and other common cyst-forming nematodes. Journal of Nematology 19, 541.
  • Mulholland V, Carde L, O’Donnell KJ, Fleming CC & Powers TO (1996) Use of the polymerase chain reaction to discriminate potato cyst nematode at the species level. In: Diagnostics in Crop Protection (Ed. MarshallG), pp. 247252. British Crop Protection Council, Farnham (GB).
  • OEPP/EPPO (1991) EPPO Standards PM 3/30 phytosanitary procedures for Globodera pallida & G. rostochiensis. Soil sampling methods. Bulletin OEPP/EPPO Bulletin 21, 233240.
  • OEPP/EPPO (2006) EPPO Standards PM 3/68 Testing of potato varieties to assess resistance to Globodera rostochiensis and Globodera pallida. EPPO Bulletin 36(3), 419420.
  • Phillips MS, Forrest JMS & Wilson LA (1980) Screening for resistance to potato cyst nematode using closed containers. Annals of Applied Biology 96, 317322.
  • Sambrook J, Fritsch EF & Maniatis T (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY (US).
  • Schots A, Gommers FJ & Egberts E (1992) Quantitative ELISA for the detection of potato cyst nematodes in soil samples. Fundamental and Applied Nematology 15, 5561.
  • Shields R, Fleming CC & Stratford R (1996) Identification of potato cyst nematodes using the polymerase chain reaction. Fundamental and Applied Nematology 19, 167173.
  • Siddiqi MR (2000) Key to genera of Heteroderinae. In: Tylenchida: Parasites of Plants and Insects (Ed SiddiqiMR), pp. 395396. CABI Publishing, Wallingford, Oxon (GB).
  • Stone AR (1972) Heterodera pallida n. sp. (Nematoda: Heteroderidae) a second species of potato cyst nematode. Nematologia 21, 8188.
  • Stone AR (1973a) CIH Descriptions of Plant-Parasitic Nematodes No 16 Globodera rostochiensis. CAB International, Wallingford (GB).
  • Stone AR (1973b) CIH Descriptions of Plant-Parasitic Nematodes No 15 Globodera pallida. CAB International, Wallingford (GB).
  • Thiéry M & Mugniéry D (1996) Interspecific rDNA restriction fragment length polymorphism in Globodera species, parasites of solanaceous plants. Fundamental and Applied Nematology 19, 471479.
  • Turner SJ (1998) Sample preparation, soil extraction and laboratory facilities for the detection of potato cyst nematodes. In: Potato Cyst Nematodes: Biology, Distribution and Control (Ed. MarksRJ & BrodieBB), pp. 7590. CAB International, Wallingford (GB).
  • Van Riel HR & Mulder A (1998) Potato cyst nematodes (Globodera species) in western Europe. In: Potato Cyst Nematodes: Biology, Distribution and Control (Ed. MarksRJ & BrodieBB), pp. 271298. CAB International, Wallingford (GB).
  • Vrain TS, Wakarchuk DA, Levesque AC & Hamilton RI (1992) Interspecific rDNA restriction fragment length polymorphism in the Xiphinema americanum group. Fundamental and Applied Nematology 16, 563573.
  • Winfield AL, Enfield MA & Foreman JH (1987) A column elutriator for extracting cyst nematodes and other small invertebrates from soil samples. Annual Applied Biology 111, 223231.
  • Wouts WM & Baldwin JG (1998) Taxonomy and identification. In: The Cyst Nematodes (Ed. SharmaSB), pp. 83122. Kluwer, Dordrecht (NL).
  • Zouhar M, Rysanek O & Kocova M (2000) Detection and differentiation of the potato cyst nematodes Globodera rostochiensis and Globodera pallida by PCR. Plant Protection Science 36, 8184.

Appendices

  1. Top of page
  2. Abstract
  3. Introduction
  4. Identity
  5. Detection
  6. Identification
  7. Reference material
  8. Reporting and documentation
  9. Further information
  10. Acknowledgements
  11. References
  12. Appendices

Appendix 1 – Methods for extracting cysts of Globodera spp. from soil

The methods described here involve basic extraction techniques. Laboratories may improve them (e.g. automatisation).

If viability tests have to be performed, methods that use heat to dry samples are not recommended (samples should not be dried at a higher temperature than 25°C and not lower than 40% air humidity).

Methods for cyst extraction based on flotation techniques
Flask and paper-strip methods

These methods are based on the characteristic that dried cysts float. The dried soil sample is added to a beaker, flask or white dish that is filled with water. The suspension is well stirred. After 30 s to some minutes, depending on the soil type, the water is cleared and the liquid will only contain the floating organic debris and cysts. Adding a drop of detergent will cause the cysts to move to the edge and the cysts can be picked out by hand using a brush. Other ways are carefully decanting, or using a paper strip around the beaker and raising the water level so cysts can adhere to it. A variety of methods to isolate the cysts from the debris are in use (Turner, 1998).

Fenwick can

The Fenwick can is an apparatus that has been in use for many years (Fig. 8). The apparatus is a container, tapering towards the top, with a sloping collar around the outside of the rim which collects overflow and directs it towards an outlet. The can has a sloping internal base with a drain plug at its lowest point. The can is first filled with water and the soil sample is added by washing through a 1-mm sieve supported on a long-stemmed funnel above the can. The organic matter will immediately rise and overflow onto the collar and be collected on two sieves with an aperture of 840 and 250 μm. Most of the cysts in the soil sample will be collected at this stage. The funnel above the can is then removed and the soil at the base of the can is elutriated by means of water flowing rapidly through a long glass or metal tube. The tube is inserted deep into the can to stir the sediment and release any trapped cysts; this is continued for approximately 1 min. The cysts are collected on the 250-μm aperture sieve for further processing.

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Figure 8.  Vertical-section diagram of Fenwick can.

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Some automated versions of this apparatus exist (e.g. carousel).

Schuiling centrifuge

The Schuiling centrifuge is a semi-automatic extraction method. The air-dried, 200 mL soil sample is added to a transparent cylindrical container half-filled with water. The contents are swirled with a rotating two-pronged fork at 450–500 rev min−1, creating a vortex and causing cysts and similar-sized floating particles to be forced to the centre through a wire-mesh cylinder (1.5 mm aperture). The mesh cylinder is fixed above a tube of the same diameter leading down to an outlet. While swirling, more water is added around the inside of the main container washing off any adhering debris and cysts which are channelled to the outlet with the rest. The apparatus cleans itself after each sample processing. Further separation of cysts is by a special cleaning process involving the so-called Schuiling can and special sieves. In some laboratories the Schuiling units have been modified to suit different soils and conditions: the modifications include additional spinning and cleaning time, larger collecting sieves, an improved plastic cleaning ‘can’ for reducing the amount of debris, and removal of the electrical parts from the apparatus to the wall above for safety reasons. Cysts are collected on a 250-μm aperture sieve for further processing.

Methods for cysts extraction based on elutriation techniques

These methods are based on the difference in density of cyst in comparison to soil particles and can be used for wet soil. At the base of a (conical) column water enters through a perforated tube at a constant rate (minimal 0.6 L min−1). Soil is added into the column using a funnel. A small plate baffles the outlet of the funnel so that soil does not fall down the column too quickly. Water containing flows from the overflows spout and are collected on a pair of sieves (250 μm aperture).

Alternatively, the suspension containing the cysts and organic debris can be carefully decanted to separate from the test of the soil contents, or a paper strip can be attached around the internal rim of the beaker so that when the water level is raised, the cysts can adhere to it.

Seinhorst elutriator

The equipment is to be used for extraction of cysts both from dry and wet soil samples (Fig. 9).

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Figure 9.  (A) Seinhorst elutriator. (B) Cross section of the base of the Wye washer.

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The elutriator is first filled with water and the constant upward water stream entering the base of elutriator is opened. This stream allows the ‘wet’ heavy cysts to float in the suspension. Next the wet soil sample is placed on the upper sieve (1 mm pore). The sample is washed into elutriator by moving the sieve up and down. Heavy particles (sand, etc.) will settle in the bottom part of the elutriator. The cysts and some (light) debris are caught on 0.25 mm pore sieve or a bucket with such a sieve on the bottom. Fine particles pass the sieve. The sample is run 2–5 min depending on soil type. After washing is finished the constant upward water stream entering the base of elutriator is closed and the content of the apparatus is drawn off through the lower outlet onto a sieve or bucket. The collected material is destined for further processing.

Some automated versions of this apparatus exist.

Wye washer

The apparatus (Winfield et al., 1987) is constructed of a 50-cm length of 15-cm diameter clear acrylic tube which, at its lower end, is held inside two tight-fitting concentric PVC sleeves (Fig. 9). Water enters through an inlet pipe on the outer sleeve and is caused to swirl by means of an arrangement of grooves and angled holes on the inner sleeve and the acrylic tube. At the top of the tube is a spout which directs overflow onto sieves of similar size to those used with the Fenwick can (i.e. aperture of 840 and 250 μm). A soil sample, up to 1000 mL is added to a small quantity of water in the Wye washer. More water is added as rapidly as possible, to break up the soil until the rim is reached, whereupon the flow is briefly stopped and then increased gradually to about 10 L min−1 for 10 min. The overflow carries: (1) small soil particles, which will pass through both sieves; (2) large organic debris which will be retained by the upper larger sieve; (3) cysts and similar sized organic particles, collected on the 250 μm aperture sieve.

Appendix 2 – Bioassays

Test A Bioassay (method performed in Germany and Austria).

This method relies on the principle that if potato cyst nematodes are present in a soil sample (even in very low numbers) they will multiply when given access to the roots of growing potato plantlets in a small container. The presence of developing cysts on the roots can then be observed through the transparent walls of the special containers used4.

Depending on the size of the container about 100–200 mL of soil should be put into each container, ensuring that the soil remains suitably moist. Eyes are cut from well chitted, certified tubers with a circular blade (diameter approx. 3 cm) and placed in the containers. Bioassay in autumn/winter requires chitting of tubers (through fumigation or treatment with gibberellic acid). To avoid growth of fungi eye cuttings should be let to dry for half a day at room temperature before they are placed on the soil samples in the containers (eyes upwards) and covered with nematode-free soil. Control containers with known infestations are used in each test.

Containers are placed close together on a planting table shading each other to prevent the growth of algae on the transparent walls. To provide for an optimal host-parasite interaction air temperature in the glasshouse is ideally maintained at 16/22°C (night/day) and always kept below 25°C (possibly giving additional light in winter). Containers should be watered moderately to achieve an optimal root penetration of the soil. Watering may be done manually or by trickle irrigation. Surplus irrigation water can run off through a hole in the bottom of the containers. The risk of contaminating healthy samples by means of adjacent infested samples has been shown to be unlikely. It might be necessary to take measures against foliar blight during the course of the bioassay. If an individual plant should die, the soil in the container should be tested for cysts using the Fenwick can or a related method.

Visual observation of females and cysts is done when cysts are observed in the control containers, generally after 6–10 weeks of cultivation. Before counting of females and cysts, potato leaves are cut with pruning shears. New cysts are visible on the roots through the transparent walls of the containers, when infection levels are high. To detect low levels of infestation, it is advised to inspect roots and soil after removing from the container and to extract cysts from the soil when no infection is detected visually.

This can also be performed in closed containers kept in a dark room (Phillips et al., 1980).

Test B Test of reproduction (method performed in Norway).

The infective success of PCN is tested on potato plants in 500 mL pots with sand, using nylon bags with cysts (up to 20) as inoculation units. It is recommended to treat tubers with gibberellic acid in order to induce and synchronize the germination. Each pot is filled with 1/3 of the total soil volume, supplied with one potato, and then filled up with sand. Each pot receive a nylon bag with cysts and the pots are placed in a randomised fashion in a growth cabinet with approximate day/night temperatures of 20°C/16°C and 18 h light period. The pots should receive mineral nutrients and water as required. After 3 months the shoots are cut and the soil and roots are air-dried. The newly formed cysts are extracted from soil (for instance by the Fenwick can), and collected and counted. Each new cyst represents a successful infection and hence a measure of the infection potential of the population.

Appendix 3 – Molecular tests

The following PCR tests are recommended for the identification of G. rostochiensis and G. pallida by use of species-specific primers:

  • • 
    A commercially available test developed by Blgg (http://www.blgg.nl), a real-time SYBR-green assay based on rDNA (LSU) sequences based PCR (D).

The PCR test of Bulman & Marshall was evaluated using 269 field samples containing PCN, and proved more sensitive than ELISA and Isoelectric focusing (IEF) (Ibrahim et al., 2001).

For each method the DNA isolation methods are described in detail as in reference cited. There are also other extraction methods available (see Fleming et al., 2000). Alternatively, commercial kits for DNA isolation are available.

(A) Multiplex PCR test (Bulman & Marshall, 1997)
1. General information
1.1. Test developed by Bulman & Marshall (1997). The test can only be used on nematodes morphologically identified as Globodera sp., as the primers are not specific for Globodera spp.
1.2. Different G. pallida and G. rostochiensis populations, from different pathotype and geographical origin were used. Full cysts are nucleic acid source.
1.3. The assay is designed to the 18S rRNA gene and the internal transcribed spacer ITS1 region.
1.4. Oligonucleotides: One universal primer ITS5 (5′-GGAAGTAAAAGTCGTAACAAGG-3′), and also specific primers for G. pallida (PITSp4: 5′-ACAACAGCAATCGTCGAG-3′) and G. rostochiensis PITSr3: 5′-AGCGCAGACATGCCGCAA-3′ result in amplicons of 265 bp for G. pallida and 434 bp for G. rostochiensis. The primers can be used in a mixture for a multiplex PCR.
1.5. Taq DNA polymerase (Life technologies, Carlsbad, CA, US) used for the amplification.
1.6. Nucleotides are used at a final concentration of 0.16 mM each.
1.7. Molecular grade water (MGW) is used to make up reaction mixes.
2. Methods
2.1. Nucleic acid Extraction and Purification
Twenty cysts were ground in Eppendorf tubes, using plastic micro-pestles (Treff, Degersheim, CH), with 200 μL of solution containing 5 M guanidine isothiocyanate, 10 mM EDTA, 50 mM Tris–HCl (pH 7.5), and 8% mercaptoethanol. After room temperature incubation for up to 1 h, the DNA containing solution was extracted once with equal volumes of phenol and chloroform-isoamyl alcohol (24:1) and once with chloroform–isoamyl alcohol then precipitated with 0.3 M sodium acetate and two volumes ethanol. DNA was resuspended in 100 μL of H2O.
2.2. Polymerase Chain reaction
2.2.1. Master mix (concentration per 25 μL single reaction)
2 mM MgCl2
0.16 mM each dNTP
250 μM each primer is mentioned in the original publication but laboratories performing the test use a final concentration ranging from 0.15 to 1 μM
0.6 U Taq DNA Polymerase (Life Technologies)
2.2.2. PCR cycling parameters
2 min 94°C, 35 cycles of 30 s 94°C, 30 s 55°C, 2 min 72°C, final elongation 5 min 72°C
3 Essential Procedural Information
3.1. Analysis of DNA fragments:
DNA fragments are separated by electrophoresis on agarose gel and visualized under UV light according to standard procedures (e.g. Sambrook et al., 1989)
3.2. Identification of species: expected amplicons for G. pallida: 265 bp and for G. rostochiensis 434 bp.
3.3. A negative control (no DNA target) should be included in every experiment to test for contamination as well as a positive control (DNA from a reference strain of the pathogen).
(B) ITS PCR-RFLP test (Fleming et al., 2000)
1. General information
1.1. Test developed by Fleming et al. (2000). The ITS1 rDNA region is the target region. The test can only be used on nematodes morphologically identified as Globodera sp., as the primers are not specific for Globodera spp.
1.2. Full cysts are nucleic acid source. A single cyst can be used.
1.3. Oligonucleotides: Primers rDNA2 5-TTG ATT ACG TCC CTG CCC TTT-3 and rDNA1 5.8 s 5-ACGAGCCGAGTGATCCACCG-3 result in a 750-bp amplicon for Globodera species.
1.4. Taq DNA polymerase used for amplification and enzymes AluI and HinfI for amplicon restriction.
1.5. Molecular grade water (MGW) is used to make up reaction mixes.
2. Methods
2.1. Nucleic acid Extraction and Purification
DNA is extracted from cysts by standard procedures. Alternatively, crude DNA extract can be prepared by crushing or boiling a juvenile nematode in 10 μL of water and then used directly in a PCR reaction.
2.2. Polymerase Chain reaction
2.2.1. Master mix (concentration per 25-μL single reaction)
2.5 μL Mg-free 10× reaction buffer
1.5 mM MgCl2
0.64 mM each dNTP
0.8 mM each primer
1 U Taq DNA Polymerase
1 μL target DNA
2.2.2. PCR cycling parameters
35 cycles of 45 s 94°C, 1 min 50°C, 1 min 72°C.
2.3. Restriction of PCR amplicon
2.3.1. Restriction mix (concentration per 20-μL reaction)
According to supplier’s conditions with 5 μL of PCR product and 2 units of restriction enzymes, 2 μL 10× restriction enzyme buffer.
2.3.2. Incubation time/temperature for digestion: 37°C for 4 h. It may occur that PCR product do not digest to completion. In such case overnight incubation or purification of PCR products before restriction step should be performed using PCR purification kits.
3 Essential Procedural Information
3.1. Analysis of RFLP fragments:
RFLP fragments are separated by electrophoresis on 2% agarose gel in 1× TAE buffer and visualized under UV light according to standard procedures (e.g. Sambrook et al., 1989).
3.2. RFLP sizes for G. rostochiensis, G. pallida and G. tabacum are given in Table 2
Table 2.   Sizes of RFLP fragments
EnzymeG. rostochiensisG. pallidaG. tabacum
AluI 360, 160, 150, 80510, 160, 80510, 160, 80
HinfI520, 230370, 230, 150520, 230
(C) ITS PCR-RFLP test (Thiéry & Mugniéry, 1996)
1. General information
1.1. Test developed by Thiéry & Mugniéry (1996). The method can only be used on nematodes morphologically identified as Globodera sp., as the primers are not specific for Globodera spp.
1.2. Full cysts or females are nucleic acid source. G. pallida and G. rostochiensis populations from various sources (e.g. pathotypes, geographical origin) were tested. G. ‘mexicana’, G. tabacum tabacum, G. tabacum virginae and G. tabacum solanacearum were also included in this test.
1.3. Oligonucleotides: Primers 18S and 26S described by Vrain et al. (1992), 18S primer 5′-TTG ATT ACG TCC CTG CCC TTT-3′ and 26S primer 5′-GGA ATC ATT GCC GCT CAC TTT-3′ result in a 1200-bp amplicon.
1.4. Taq DNA polymerase (MP biomedicals, Illkirch, FR ex Appligene Oncor) used for amplification and enzyme Bsh1236I for amplicon restriction (other restriction enzymes were tested, but this single one is specific enough to separate the species).
1.5. Nucleotides used at a final concentration of 100 μM.
1.6. Molecular grade water (MGW) is used to make up reaction mixes; this should be purified (deionised or distilled), sterile (autoclaved or 0.45 μm filtered) and nuclease free.
1.7. Amplification is performed in a thermal cycler, e.g. Perkin-Elmer Cetius DNA Thermal Cycler 480.
2. Methods
2.1. Nucleic acid Extraction and Purification
Genomic DNA was isolated according to Folkertsma et al. (1994). An extraction with 6 M NaCl solution was done to precipitate the majority of proteins. RNA was removed by incubation with 20 μg of RNaseA at 37°C for 30 min.
2.2. Polymerase Chain reaction
2.2.1. Master mix (concentration per 50-μL single reaction)
2 mM MgCl2
100 μM each dNTP
0.5 μM each primer
0.5 U Taq DNA Polymerase (MP Biomedicals, ex Appligene Oncor)
2.2.2. PCR cycling parameters
30 cycles of 1 min 94°C, 50 s 60°C, 1 min 72°C.
2.3. Restriction of PCR amplicon
2.3.1. Restriction mix
According to supplier’s conditions.
Incubation time/temperature for digestion: overnight at the recommended temperature (see supplier’s information).
3 Essential Procedural Information
3.1. Analysis of DNA fragments:
DNA fragments are separated by electrophoresis on agarose gel (1.5% for RFLP) and visualized under UV light according to standard procedures (e.g. Sambrook et al., 1989).
3.2. Species specific RFLP patterns for G. pallida and G. rostochiensis are given in Table 3.
3.3. A negative control (no DNA target) should be included in every experiment to test for contamination as well as a positive control (DNA from a reference strain of the pathogen).
Table 3.   Restricted fragments sizes in bp (Thiéry & Mugniéry, 1996)
SpeciesBsh1236I RFLP pattern
G. rostochiensis (European populations)900, 190, 110
G. pallida (European populations)500, 400, 350, 190, 110
G. ‘mexicana’500, 400, 190, 110
G. tabacum tabacum445, 400, 190, 110
G. tabacum virginae445, 400, 190, 110
G. tabacum solanacearum445, 400, 190, 110
(D) Real-time SYBR-green assay based on rDNA (LSU) sequences
1. General information
1.1. This protocol was developed by Blgg bv. and the Laboratory of Nematology (Wageningen University, The Netherlands). The development of these species-specific LSU rDNA primers was a spin-off of a phylum-wide phylogenetic study among nematodes (based on SSU rDNA) conducted by Holterman et al. (2006). The Globodera primer sequences (from private and public databases) were aligned with 400 nematode sequences in a LSU rDNA sequence database, including 22 rDNA sequences of 5 different Globodera species (G. pallida, G. rostochiensis, G. tabacum, G. achilleae and G. artemisiae). The primers for G. pallida and G. rostochiensis only match with the LSU rDNA sequences of G. pallida and G. rostochiensis, respectively.
1.2. Determination of analytical sensitivity and analytical specificity was performed on a Biorad MyiQ real-time PCR detection system. The analytical specificity of the G. pallida and G. rostochiensis primer sets was tested with plasmid DNA against G. pallida and G. rostochiensis, respectively (simplex reaction), besides G. tabacum, G. achilleae and G. artemisiae. No false positive signals are obtained with these species. The analytical sensitivity (detection limit) is one single PCN juvenile or egg, against a background of 1000 non-target PCN juveniles or eggs. A DNA extract of well lysed cysts (juveniles or eggs) is needed as starting material. The simplex amplifications can be performed in any real-time PCR instrument. The melting curve temperature are 85.5 and 86°C for G. pallida and G. rostochiensis, respectively. Alternatively, the primers can be used for conventional PCR.
2. Methods
2.1. The LSU (28S) detection kit includes primers to be used for (real-time) PCR detection of G. pallida and G. rostochiensis.
3. Essential Procedural information
3.1. A negative and/or blank control (no DNA target) should be included in every experiment to test for contamination as well as a positive control (DNA from a reference strain of the pathogen).
3.2. More information, primer sets and contact details can be obtained from Blgg bv in The Netherlands (at http://www.blgg.com).

Appendix 4 – Visual determination5

Live eggs (Figs 10 and 11)Dead eggs (Figs 15 and 16)
aWhole egg is intactaEgg may be damaged/broken and empty
bEgg shell is smoothbEgg shell often not smooth
cEgg is clear/transparent with distinct contents or a dark line down the middle of the egg cContents have black/grey granular appearance with no structure
dCurled juvenile fill up against the egg shelldShrivelled disintegrated juvenile in egg
eSometimes clear lip region and stylet presenteNo clear lip region or stylet present
Live juveniles (Figs 12–14)Dead juveniles (Figs 17–19)
aJuvenile has clear lip region, stylet visibleaNo clear lip region, partly or completely grey/black structure
bJuvenile has strong smooth cuticlebCuticle shrivelled or not intact
cIntestine is filled with grey granular structure, solidcTransparent, body with clear patches or completely transparent
dClear lopsided distinction between pharynx and intestinedNo clear lopsided distinction between pharynx and intestine
  eJuvenile sharply bent at an angle or lying in half circle
Included in counts: HeadsNot included in counts: Tails, empty shells
image

Figure 10–19.  Characteristics of live and dead eggs and juveniles of the potato cyst nematodes.

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Appendix 5 – Hatching test

(A) Hatching test (method performed in Norway)

The hatching medium, potato root diffusate (PRD), is obtained by passing 1000 mL of tap water through a 500-mL pot with a 3-week-old potato plant growing in sand. After filtering, the PRD is stored at +3°C in the dark until needed. The PRD is used without dilution in closed glass vials (Ø = 23.5 mm; height 34 mm) functioning as hatching units. Each vial contains one cyst bag made of nylon net with 20 cysts, which is completely covered by PRD. Cysts collected in the autumn need to be exposed +4°C for 2–3 weeks for the contents to hatch. The test normally lasts for 8 weeks, but the hatch is often already high within 2 weeks. Each week the cyst bags are transferred to new hatching units with fresh PRD. The number of hatched juveniles is counted weekly and accumulated to form the total hatch. At the end of the test the juveniles remaining in the cysts are counted and the hatching expressed as a percentage of the cyst content (Fig. 20).

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Figure 20.  Cumulative hatch of Globodera rostochiensis in potato root diffusate from 3 weeks-old plants (n = 4).

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(B) Hatching test (methods performed in Sweden)

Prior to the exposition to the hatching stimulus, cysts that have been stored dry should be presoaked in water in Petri dishes, staining blocks or other suitable containers for 4–5 days. Whereas non-hydrated cysts trend to float, hydrated once sink to the bottom of the Petri dish which gives a good indication of cyst’s hydration level. During this hydration phase, the water in the containers is renewed daily to prevent bacterial and fungal growth. Repeated up and down pipetting of the cysts helps to get rid of the fungi and favours the hydration.

Potato plants for production of the hatching medium, PRD, are grown in small (200 mL) clay pots with silver sand substrate in a greenhouse. Three weeks old plants are removed from the substrate and their roots are rinsed in water after which the plants are transferred one by one into a 200-mL beakers filled with tap water and incubated at room temperature under aeration by an aquarium air pump (only the root system is immersed in the water).

After 24 h diffusate is collected, filtered and ready to use. Hydrated cysts (up to 100, depending on the number of cysts found in the sample) are soaked in the undiluted PRD. The PRD is replaced daily with a fresh one.

The test lasts until juveniles start to hatch or in total for 8 weeks.

(C) Hatching test (methods performed in France)

This test is shorter and follows the procedure described in test B with the following modifications: potato root diffusate production: sprout tubers are placed on a funnel put on plastic (transparent) beaker filled up with tap water. This assemblage is put at room temperature (around 18–19°C) in the dark for 4 weeks. The water with PRD is filtered, divided in aliquot parts and frozen until use (−20°C). After 48 h freezing time, the PRD is evaluated with the previous batch of PRD (previous production) and a reference G. pallida and G. rostochiensis populations. If the test is satisfactory, the PRD can be used for the hatching test.

Hatching test: instead of being rehydrated, cysts are deposited in a fine sieve (250 μm), placed on a small dish filled with PRD. One sieve is prepared per sample to be tested. 20 cysts of G. pallida and G. rostochiensis are put respectively in two sieves as positive controls. All samples and controls are left at room temperature in the dark until they are checked for hatching. Each sample and the controls are checked for hatching every 10 days. If juveniles have hatched, the test is considered positive and the result is that the cysts are viable. If no juvenile has hatched after 10 and 20 days the PRD is removed and new PRD is added (from the validated batch). The test is considered negative if no hatching has occurred within 30 days provided that the positive controls have hatched. When positive controls do not hatch, the viability tests are not considered valid.