Globodera rostochiensis (yellow potato cyst nematode)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Latitude/Altitude Ranges
- Air Temperature
- Natural enemies
- Means of Movement and Dispersal
- Pathway Causes
- Pathway Vectors
- Plant Trade
- Impact Summary
- Economic Impact
- Risk and Impact Factors
- Uses List
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Globodera rostochiensis (Wollenweber, 1923) Skarbilovich, 1959
Preferred Common Name
- yellow potato cyst nematode
Other Scientific Names
- Globodera rostochiensis (Wollenweber, 1923) Behrens, 1975
- Heterodera (Globodera) rostochiensis Wollenweber, 1923 (Skarbilovich, 1959)
- Heterodera rostochiensis Wollenweber, 1923
- Heterodera schachtii rostochiensis Wollenweber, 1923
- Heterodera schachtii solani Zimmerman, 1927
International Common Names
- English: eelworm, golden; eelworm, potato root; golden eelworm; golden nematode; golden nematode eelworm; golden nematode of potato; golden potato cyst nematode; nematode of potato, golden; potato cyst nematode; potato golden nematode; potato root eelworm
- Spanish: nematodo dorado; nematodo dorado de la papa
- French: anguillule a kyste de la pomme de terre; anguillule des racines de la pomme de terre; nématode doré; nématode doré de la pomme de terre
Local Common Names
- Denmark: kartoffelal; kartottelcystenematod
- Finland: peruna-ankeroinen
- Germany: Aelchen, Goldfarbenes Kartoffelzysten-; Aelchen, Kartoffel-; Nematode, Kartoffel-
- Iran: nematode sibsamini
- Italy: Anguillula della patata
- Netherlands: Aardappelcystenaaltje
- Norway: potetcystenematode
- Sweden: potatiscystnematod
- HETDRO (Globodera rostochiensis)
Summary of InvasivenessTop of page
G. rostochiensis is a world wide pest of temperate areas, including both temperate countries and temperate regions of tropical countries, for example India’s Nigrilis region. Distribution is linked to that of the potato crop. Potato cyst nematode is considered to have originated from the Andes region of South America, from where it spread to Europe with potatoes. The ease with which it has been transported across continents proves what a resilient pest it is. The cyst form which adheres to host roots, stolons and tubers and to soil particles during transportation gives rise to new infestations where climate and food source are both available and favourable.
Secondary means of dispersal is through the movement of contaminated farm machinery, farming implements and contaminated footwear. Cysts are also successfully spread by wind dispersal, during winter storms or sand storms where top soil is redistributed. Rain which causes flooding and water to run off fields into trenches or irrigation channels also redistributes cysts into adjoining areas.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Nematoda
- Class: Secernentea
- Order: Tylenchida
- Family: Heteroderidae
- Genus: Globodera
- Species: Globodera rostochiensis
Notes on Taxonomy and NomenclatureTop of page
G. rostochiensis was first described by Wollenweber in 1923 as Heterodera rostochiensis. The type locality is Tessin, Mecklenburg, near Rostock, Germany (Golden and Ellington, 1972) and the host neotype is Solanum tuberosum. The species should not be confused with the very rare Globodera leptonepia (Cobb and Taylor, 1953). The origin of G. rostochiensis is considered to be the Andes mountains in South America (Krall and Krall, 1978), from where it was introduced into Europe in contaminated soil adhering to potato tubers. In 1959, Skarbilovich erected the subgenus Globodera to accommodate the round cyst nematode species of which there are several, including the potato cyst nematodes. Behrens (1975) raised Globodera to generic level, before Mulvey and Stone (1976).
In 1973, Stone described a second species of potato cyst nematode, Heterodera pallida, and it is worthy of note that information concerning H. rostochiensis before this date would have involved both species.
DescriptionTop of page
The eggs of G. rostochiensis are always retained within the cyst body and no egg sacs are produced. The eggshell surface is smooth and no microvilli are present.
Length=101-104 µm; width="46"-48 µm; L/W ratio=2.1-2.5
The females emerge from the root cortex about one month to six weeks after invasion by the second-stage juveniles. They are pure white initially, turning golden yellow on maturation. Mature females are approximately 500 µm in circumference without a cone. The cuticle of the female sometimes has a thin subcrystalline layer.
Stylet length=23 µm ± 1 µm; stylet base to dorsal oesophageal gland duct=6 µm ± 1 µm; head width at the base=5.2 µm ± 0.7 µm; head tip to median bulb=73 µm ± 14.6 µm; median bulb valve to excretory pore=65 µm ± 2.0 µm; head tip to excretory pore=145 µm ± 17 µm; mean diameter of the median bulb=30µm ± 3.0 µm; mean diameter of the vulval basin=22 µm ± 2.8 µm; vulval slit length=9.7 µm ± 2.0 µm; anus to vulval basin=60 µm ± 10 µm; number of cuticular ridges between the anus and vulva=21 ± 3.0.
The female head bears one to two annules and the neck region has numerous tubercules, which can be seen using a scanning electron microscope. The head skeleton is hexaradiate and weak. The stylet is divided equally in length between the conus and the shaft. An important diagnostic feature is the backward slope of the stylet knobs. The median bulb is large and circular and well developed. The large paired ovaries often displace the oesophageal glands. The excretory pore is well defined at the base of the neck. The posterior of the female, at the opposite pole to the neck and head, is referred to as the vulval basin and is contained within a rounded depression. The vulval slit is located in the centre of this region flanked on either side by papillae, which usually fill the translucent areas of cuticle in crescentic shape, from the slit to the edge of the fenestra. The anus is distinct and is often seen at the point in the cuticle where the 'V' shape tapers to an end. The number of cuticular ridges found in the area between the anus and the edge of the fenestra is counted as an aid to identification of Globodera species. The entire cuticle is covered in small subsurface punctations.
Length without neck=445 µm ± 50 µm; width="382" µm ± 60 µm; neck length=104 µm ± 19 µm; mean fenestral diameter=19.0 µm ± 2.0 µm; anus to fenestra=66.5 µm ± 10.3 µm; Granek's ratio=3.6 ± 0.8.
Cysts contain the eggs, the progeny for the next generation, and are formed from the hardened dead cuticle of the female. Newly produced cysts may still show an intact vulval basin but older cysts, particularly those which have been in the soil for many seasons, will have lost all signs of their genitalia with only a hole in the cuticle to show the position of the fenestral basin.
Length=0.89 -1.27 mm; width at excretory pore=28 µm ± 1.7 µm; head width at base=11.8 µm ± 0.6 µm; head length=7.0 µm ± 0.3 µm; stylet length=26 µm ± 1.0 µm; stylet base to dorsal oesophageal gland duct=5.3 µm ± 1.0 µm; head tip to median bulb valve=98.5 µm ± 7.4 µm; median bulb valve to excretory pore=74 µm ± 9µm; head tip to excretory pore=172 µm ± 12.0 µm; tail length=5.4 µm ± 1.0 µm; tail width at anus=13.5 µm ± 0.4 µm; spicule length=35.0 µm ± 3.0 µm; gubernaculum length=10.3 µm ± 1.5µm.
The male is vermiform in shape with a short tail and no bursa. On fixation, the body assumes a curved shape with the posterior region twisted at a 90 degree angle to the remainder of the body. There are four incisures in the mid-body i.e. three bands which terminate on the tail. The rounded head is offset and bears 6-7 annules. The head is strongly developed having a hexaradiate skeleton. The cephalids are located at body annules 2-4 and 6-9, respectively. The stylet is strong and has backward sloping knobs. The median bulb is well developed and has a large crescentic valve. The nerve ring is located around the oesophagus between the median bulb and the intestine. The hemizonid is found 2-3 annules anterior to the excretory pore and is itself two annules in length. The hemizonion is approximately nine body annules posterior to the excretory pore and is one annule in length. The single testis fills half the body cavity. The paired spicules are arcuate and end with single tips. The gubernaculum is around 10 µm in length and 2 µm in thickness and lies in a position dorsal to the spicules.
Body length=468 µm ± 100 µm; width at excretory pore=18 µm ± 0.6 µm; head length=4.6µm ± 0.6 µm; stylet length=22 µm ± 0.7 µm; head tip to median bulb valve=69 µm ± 2.0 µm; median bulb valve to excretory pore=31 µm ± 2.0 µm; head tip to excretory pore=100 µm ± 2.0 µm; tail length=44 µm ± 12 µm; tail width at anus=11.4 µm ± 0.6 µm; hyaline tail length=26.5 µm ± 2.0 µm.
The second-stage juvenile hatches from the egg, the first moult taking place within the egg. The juvenile, like the male, is vermiform with a rounded head and finely tapered tail. The hyaline portion of the tail represents about two thirds of its length. The lateral field has four incisures in the mid-body region reducing to three at the tail terminus and anterior end. The head is slightly offset and bears four to six annules. The head skeleton is well developed and hexaradiate in form. The cephalids are located at body annules 2-3 and 6-8, respectively. The stylet is strong, the conus being about 45% of the total length. The stylet knobs are an important diagnostic feature and typically slope backwards. The median bulb is well developed and elliptical in shape, having a large central valve. The nerve ring encircles the oesophagus between the median valve and the intestine. The hemizonion is about one body annule in width and is located five body annules posterior from the excretory pore. The hemizonid is around two body annules in width and is found just anterior to the excretory pore. The gonad primordium is four-celled and located at around 60% of the body length.
Other measurements can be found in Granek (1955), Spears (1968), Green (1971), Greet (1972), Golden and Ellington (1972), Hesling (1973, 1974), Mulvey (1973), Behrens (1975), Mulvey and Golden (1983), Othman et al. (1988) and Baldwin and Mundo-Ocampo (1991).
DistributionTop of page
G. rostochiensis is widely distributed in potato-growing regions, including the temperate regions of tropical countries.
See also CABI/EPPO (1998, No. 162).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Armenia||Present||Introduced||Invasive||Iskandaryan and Arutyunyan, 1990; CABI/EPPO, 2011; EPPO, 2014|
|India||Restricted distribution||Introduced||1961||Invasive||Prasad, 1996; CABI/EPPO, 2011; EPPO, 2014|
|-Kerala||Absent, reported but not confirmed||CABI/EPPO, 2011; EPPO, 2014|
|-Tamil Nadu||Present||Introduced||1977||Invasive||Prasad, 1996; CABI/EPPO, 2011; EPPO, 2014|
|Indonesia||Restricted distribution||2003||Indarti et al., 2004; CABI/EPPO, 2011; EPPO, 2014|
|-Java||Present||Introduced||2003||Invasive||Indarti et al., 2004; CABI/EPPO, 2011; EPPO, 2014|
|Iran||Present||Gitty and Maafi, 2010; CABI/EPPO, 2011; EPPO, 2014|
|Israel||Eradicated||1954||CABI/EPPO, 2000; CABI/EPPO, 2011; EPPO, 2014|
|Japan||Restricted distribution||Introduced||1972||Invasive||Aihara et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|-Hokkaido||Present||Introduced||1972||Invasive||Inagarki, 2004; CABI/EPPO, 2011; EPPO, 2014|
|-Kyushu||Present||Introduced||1992||Invasive||Aihara et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|Lebanon||Present||Introduced||Invasive||Ibrahim et al., 2000; CABI/EPPO, 2011; EPPO, 2014|
|Malaysia||Absent, unreliable record||EPPO, 2014|
|Oman||Present||Introduced||Invasive||Mani et al., 1993; CABI/EPPO, 2011; EPPO, 2014|
|Pakistan||Present||Introduced||1980||Invasive||Munir et al., 2004; CABI/EPPO, 2011; EPPO, 2014|
|Philippines||Present||Introduced||1983||Invasive||Davide and Zorilla, 1995; CABI/EPPO, 2011; EPPO, 2014|
|Sri Lanka||Restricted distribution||Introduced||1991||Invasive||Ekanayake, 1991; CABI/EPPO, 2011; EPPO, 2014|
|Tajikistan||Restricted distribution||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Turkey||Present, few occurrences||Introduced||Invasive||Enneli and Öztürk, 1995; CABI/EPPO, 2011; EPPO, 2014|
|Algeria||Restricted distribution||Introduced||1940s||Invasive||FRÉZAL, 1954; CABI/EPPO, 2011; EPPO, 2014|
|Egypt||Absent, unreliable record||Kleynhans, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Kenya||Present||Mwangi et al., 2015|
|Libya||Present||Introduced||1981||Invasive||Ben et al., 1981; CABI/EPPO, 2011; EPPO, 2014|
|Morocco||Absent, formerly present||CABI/EPPO, 2011; EPPO, 2014|
|Sierra Leone||Present||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|South Africa||Restricted distribution||Introduced||1950||Invasive||Kleynhans, 1998; CABI/EPPO, 2011; EPPO, 2014|
|-Canary Islands||Present||Introduced||1961||Gonzalez and Phillips, 1996; CABI/EPPO, 2011; EPPO, 2014|
|Tunisia||Restricted distribution||Introduced||1977||Invasive||Kleynhans, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Zimbabwe||Absent, intercepted only||CABI/EPPO, 2011; EPPO, 2014|
|Canada||Present, few occurrences||Introduced||1962||Invasive||IPPC, 2009; CABI/EPPO, 2011; EPPO, 2014|
|-Alberta||Present, few occurrences||CABI/EPPO, 2011; EPPO, 2014|
|-British Columbia||Present, few occurrences||Introduced||Invasive||Orchard, 1965; CABI/EPPO, 2011; EPPO, 2014|
|-Newfoundland and Labrador||Restricted distribution||Introduced||1962||Invasive||Proudfoot, 1977; CABI/EPPO, 2011; EPPO, 2014|
|-Quebec||Present, few occurrences||Introduced||Invasive||Mahran et al., 2010; CABI/EPPO, 2011; EPPO, 2014|
|Mexico||Restricted distribution||Desgarennes et al., 2009; CABI/EPPO, 2011; EPPO, 2014|
|USA||Restricted distribution||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-Delaware||Eradicated||CABI/EPPO, 2011; EPPO, 2014|
|-New York||Present, few occurrences||Introduced||1941||Invasive||NAPPO; CANNON, 1941; CABI/EPPO, 2011; EPPO, 2014|
Central America and Caribbean
|Costa Rica||Absent, formerly present||CABI/EPPO, 2011; EPPO, 2014|
|Cuba||Absent, intercepted only||CABI/EPPO, 2011|
|Panama||Present||Introduced||1968||Invasive||Brodie, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Argentina||Absent, formerly present||1956||Franco et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|Bolivia||Present||Native||Bendezu et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|Brazil||Absent, invalid record||CABI/EPPO, 2011; EPPO, 2014|
|Chile||Restricted distribution||Native||Cubillos and Fernández, 1990; CABI/EPPO, 2011; EPPO, 2014|
|Colombia||Restricted distribution||Native||BAEZA, 1972; CABI/EPPO, 2011; EPPO, 2014|
|Ecuador||Restricted distribution||Native||Franco et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|Peru||Present||Native||Picard et al., 2007; CABI/EPPO, 2011; EPPO, 2014|
|Venezuela||Present||Native||DAO et al., 1971; CABI/EPPO, 2011; EPPO, 2014|
|Albania||Restricted distribution||Introduced||1982||Invasive||Jovani, 1994; CABI/EPPO, 2011; EPPO, 2014|
|Austria||Widespread||Introduced||1940s||Invasive||Riel and Mulder, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Belarus||Present||Introduced||1957||Invasive||Ponin et al., 1978; CABI/EPPO, 2011; EPPO, 2014|
|Belgium||Restricted distribution||Introduced||1949||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Bosnia-Hercegovina||Present||Ostojic et al., 2011; EPPO, 2012; EPPO, 2014|
|Bulgaria||Restricted distribution||Introduced||1970s||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Croatia||Present, few occurrences||Introduced||Invasive||Grubisic et al., 2007; CABI/EPPO, 2011; Grubisic et al., 2013; EPPO, 2014|
|Cyprus||Restricted distribution||Introduced||1974||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Czech Republic||Restricted distribution||Introduced||1954||Invasive||Zouhar and Rysanek, 2002; CABI/EPPO, 2011; EPPO, 2014|
|Denmark||Widespread||Introduced||1928||Invasive||Hansen, 1988; CABI/EPPO, 2011; IPPC, 2013; EPPO, 2014; NPPO of Denmark, 2019|
|Estonia||Restricted distribution||Introduced||1953||Invasive||Koppel and Tsahkna, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Faroe Islands||Present||Introduced||1951||Invasive||Jakobsen, 1973; CABI/EPPO, 2011; EPPO, 2014|
|Finland||Restricted distribution||Introduced||1946||Invasive||Heikkilä and Tilikkala, 1992; CABI/EPPO, 2011; EPPO, 2014|
|France||Restricted distribution||Introduced||1948||Invasive||Riel and Mulder, 1998; CABI/EPPO, 2011; EPPO, 2014|
|-France (mainland)||Restricted distribution||CABI/EPPO, 2011|
|Germany||Widespread||Introduced||1913||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Greece||Restricted distribution||Introduced||1954||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-Crete||Present||Introduced||2004||Invasive||Tzortzakakis et al., 2004; CABI/EPPO, 2011; EPPO, 2014|
|-Greece (mainland)||Restricted distribution||Koliopanos, 1976; CABI/EPPO, 2011|
|Hungary||Restricted distribution||Introduced||1980||Invasive||Elekes-Kaminszky et al., 2004; CABI/EPPO, 2011; EPPO, 2014|
|Iceland||Widespread||Introduced||1953||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Ireland||Restricted distribution||Introduced||1922||Invasive||Riel and Mulder, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Italy||Widespread||Introduced||1961||Invasive||CABI/EPPO, 2011; EPPO, 2011; EPPO, 2014|
|-Italy (mainland)||Widespread||Greco et al., 1994; CABI/EPPO, 2011|
|-Sicily||Present||Lombardo et al., 2011|
|Latvia||Restricted distribution||Introduced||1949||Invasive||Eglitis, 1973; CABI/EPPO, 2011; EPPO, 2014|
|Liechtenstein||Widespread||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Lithuania||Restricted distribution||Introduced||1948||Invasive||Jogaite et al., 2007; CABI/EPPO, 2011; EPPO, 2014|
|Luxembourg||Restricted distribution||Introduced||1955||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Malta||Restricted distribution||Introduced||Invasive||Lamberti et al., 1987; CABI/EPPO, 2011; EPPO, 2014|
|Netherlands||Restricted distribution||Introduced||1941||Invasive||NPPO of the Netherlands, 2013; CABI/EPPO, 2011; EPPO, 2014|
|Norway||Widespread||Introduced||1955||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Poland||Restricted distribution||Introduced||1946||Invasive||Wolny, 1992; CABI/EPPO, 2011; Przetakiewicz, 2013; EPPO, 2014|
|Portugal||Restricted distribution||Introduced||1956||Invasive||Santos et al., 1995; CABI/EPPO, 2011; EPPO, 2014|
|-Azores||Absent, invalid record||CABI/EPPO, 2011; EPPO, 2014|
|-Madeira||Present||EPPO, 2009; CABI/EPPO, 2011; EPPO, 2014|
|-Portugal (mainland)||Restricted distribution||CABI/EPPO, 2011|
|Romania||Restricted distribution||Introduced||1986||Invasive||Rojancovski and Dehebanu, 1986; CABI/EPPO, 2011; EPPO, 2014|
|Russian Federation||Restricted distribution||Introduced||1945||Invasive||Subbotin et al., 1999; CABI/EPPO, 2011; EPPO, 2014|
|-Central Russia||Present||Introduced||Invasive||Gus'kova and Makovskaya, 1977; CABI/EPPO, 2011; EPPO, 2014|
|-Eastern Siberia||Present||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-Northern Russia||Present||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-Russian Far East||Present||Introduced||Invasive||Shvydkaya and Eroshenko, 1997; CABI/EPPO, 2011; EPPO, 2014|
|-Southern Russia||Present||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-Western Siberia||Present||Introduced||Invasive||Marks and Rojancovski, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Serbia||Present, few occurrences||Introduced||Invasive||Krnnjac et al., 2002; CABI/EPPO, 2011; EPPO, 2014|
|Slovakia||Restricted distribution||Introduced||Invasive||Marks and Rojancovski, 1998; CABI/EPPO, 2011; EPPO, 2014|
|Slovenia||Present, few occurrences||Introduced||Invasive||Sirca and Urek, 2004; Sirca and Urek, 2005; CABI/EPPO, 2011; EPPO, 2014|
|Spain||Restricted distribution||Introduced||Invasive||Martinez-Beringola et al., 1988; CABI/EPPO, 2011; EPPO, 2014|
|-Balearic Islands||Present, few occurrences||Introduced||Invasive||Andrés et al., 2006; CABI/EPPO, 2011; EPPO, 2014|
|-Spain (mainland)||Restricted distribution||CABI/EPPO, 2011|
|Sweden||Widespread||Introduced||1922||Invasive||Manduric and Andersson, 2003; CABI/EPPO, 2011; EPPO, 2014|
|Switzerland||Restricted distribution||Introduced||1958||Invasive||Riel and Mulder, 1998; CABI/EPPO, 2011; EPPO, 2014|
|UK||Restricted distribution||Introduced||Invasive||Minnis et al., 2002; CABI/EPPO, 2011; EPPO, 2014|
|-Channel Islands||Present||Introduced||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|-England and Wales||Restricted distribution||Minnis et al., 2002; CABI/EPPO, 2011; EPPO, 2014|
|-Northern Ireland||Restricted distribution||Introduced||1922||Invasive||Turner and Evans, 1998; CABI/EPPO, 2011; EPPO, 2014|
|-Scotland||Present||Introduced||1913||Invasive||CABI/EPPO, 2011; EPPO, 2014|
|Ukraine||Restricted distribution||Introduced||Invasive||Pylypenko et al., 2005; CABI/EPPO, 2011|
|Australia||Restricted distribution||Introduced||1986||Invasive||Stanton, 1986; IPPC, 2008; CABI/EPPO, 2011; EPPO, 2014|
|-Victoria||Restricted distribution||Introduced||1991||Invasive||Hinch et al., 1998; CABI/EPPO, 2011; EPPO, 2014|
|-Western Australia||Eradicated||Stanton, 1987; IPPC, 2010; CABI/EPPO, 2011; EPPO, 2011; EPPO, 2014; IPPC, 2015|
|New Zealand||Widespread||Introduced||1972||Invasive||CABI/EPPO, 2000; Dale, 1973; CABI/EPPO, 2011; EPPO, 2014|
|Norfolk Island||Present||Introduced||Invasive||Marshall, 1998; CABI/EPPO, 2011; EPPO, 2014|
History of Introduction and SpreadTop of page
The potato was first domesticated and cultivated by the Andean peoples as long as 8,000 years ago. When Peru was invaded by the Spanish in 1531, potatoes were already an important source of food for these peoples.
The potato was bought to Europe sometime in the sixteenth century and grown in the Seville region of Spain. An independent introduction of potato occurred in England around 1590, but not from the same source. From these introductions, potatoes spread across Europe and into other parts of the world. By the nineteenth century the potato had become more widely grown and had been dispersed over wider geographical areas by trade and the movement of populations.
With the advent of the hugely damaging potato blight (Phytophthora infestans) in the mid-nineteenth century, many people in Europe found starvation confronting them, particularly the Irish who by now were heavily dependant upon potatoes as a staple food. There are no records of potato cyst nematode damage from this time but whether it was not recognised or had not yet been introduced to areas where potato was grown is unclear. The potato now being a significant food source meant that Europeans began to search for varieties with resistance to blight from the area of origin of potatoes, South America. The new genetic material was also used to breed varieties of potato adapted to longer day lengths and with generally better adaptation to European growing conditions.
In all probability, potato cyst nematodes were introduced to Europe with the new genetic material around 1850. In 1881, Kuhn made the first record of cyst nematode damage to potato; although at the time it was described as the beet cyst nematode Heterodera schachtii. The thirty year period between 1881 and bringing in new potato material, most probably with adhering soil and cysts, was enough time for cyst nematodes to build up to damaging levels in the field and to cause crop loss.
In 1923, Wollenweber described the cysts he found on potato as the new species Heterodera rostochiensis, named after the type locality of Rostock in Germany. Later, H. pallida was described by Stone (1972) as a second species of PCN occupying an almost identical ecological niche, but having significantly different biological, morphological and biochemical characteristics.
In North America, G. rostochiensis was first discovered on Long Island, New York (Nassau County) in 1941 after a potato grower noticed isolated areas of poor plant growth (Cannon, 1941). The nematode may have come in on equipment returned from Europe after WWI - the field was an old military camp which became a potato field. The discovery led to establishment of a Federal Quarantine based on the Golden Nematode Act. Growers moved to Western New York following urbanization, and subsequent outbreaks appeared; required 8-10 years to reach a detectable level.
In the early 1970's, scientists in Mexico discovered an infestation of G. rostochiensis in the state of Guanajuato (Iverson, 1972). It was reported in South Africa in 1971 from an irrigated farm near Pretoria and then from small farms around Johannesburg and Bon Accord. Spread in Africa is detailed by Kleynhans (1998).
Two of the most recent outbreaks of Globodera rostochiensis have occurred in Quebec, Canada in 2006 (Sun et al., 2007) and in the Balearic Island of Mallorca, Spain 2006 (Andrés et al., 2006), so demonstrating the continuing need for good phytosanitary regulations and stringent quarantine measures.
IntroductionsTop of page
Risk of IntroductionTop of page
Potato cyst nematodes are A2 quarantine pests for EPPO (OEPP/EPPO, 1978, 1981, 2007). They are also of quarantine significance for APPPC and NAPPO. G. rostochiensis is also a quarantine pest for CPPC and IAPSC.
Virtually all areas within the EPPO region that grow potatoes are already contaminated with potato cyst nematodes. These areas are generally closely monitored. It is important to keep seed potato areas as free as possible of potato cyst nematodes. Domestic measures and import controls are justified as they help to reduce spread and introduction of new pathotypes into established areas. G. rostochiensis still seems to be the dominant species throughout Europe, with the exception of England, UK, where G. pallida is common.
Habitat ListTop of page
|Soil||Present, no further details||Harmful (pest or invasive)|
|Stored products||Present, no further details||Harmful (pest or invasive)|
|Terrestrial – Managed||Cultivated / agricultural land||Present, no further details||Harmful (pest or invasive)|
|Terrestrial ‑ Natural / Semi-natural||Arid regions||Present, no further details||Harmful (pest or invasive)|
|Coastal areas||Present, no further details||Harmful (pest or invasive)|
Hosts/Species AffectedTop of page
The major hosts of G. rostochiensis are restricted to the Solanaceae, in particular potato, tomato and aubergine (Ellenby, 1945, 1954; Mai, 1951, 1952; Winslow, 1954; Stelter, 1957, 1959, 1987; Roberts and Stone, 1981; Sullivan et al., 2007). A number of weeds in the Solanaceae are also hosts.
In addition to the main hosts listed, the following plants are hosts of G. rostochiensis:
Datura tatula, D. ferox, Hyoscyamus niger, Lycopersicon aureum, L. glandulosum, L. hirsutum, L. esculentum peruvianum, L. pimpinellifolium, L. pyriforme, L. racemigerum, Nicotiana acuminata, Physalis longifolia, P. philadelphica, Physochlaina orientalis, Salpiglossis sp., Saracha jaltomata, Solanum acaule, S. aethiopicum, S. ajanhuiri, S. ajuscoense, S. alandiae, S. alatum, S.americanum, S. anomalocalyx, S. antipoviczii, S. armatum, S. ascasabii, S. auriculatum, S. asperum, S. aviculare, S. berthaultii, S. blodgettii, S. boergeri, S. brevidens, S. brevimucronatum, S. bukasovii, S. bulbocastanum, S. calcense,S. calcense × S. cardenasii, S. caldasii, S. canasense, S. capsicibaccatum, S. capsicoides, S. cardiophyllum, S. carolinense, S. chacoense, S. chaucha, S. chenopodioides, S. chloropetalum, S. citrillifolium, S. coeruleiflorum, S. commersonii, S. curtilobum, S. curtipes, S. demissum, S. demissum × S. tuberosum, S. dulcamara, S. durum, S. elaeagnifolium, S. ehrenbergii, S. famatinae, S. fraxinifolium, S. fructo-tecto, S. garciae, S. gibberulosum, S. giganteum, S. gigantophyllum, S. gilo, S. glaucophyllum, S. goniocalyx, S. gourlayi, S. gracile, S. heterophyllum, S. heterodoxum, S. hirtum, S. hispidum, S. indicum, S. integrifolium, S. intrusum, S. jamesii, S. jujuyense, S. juzepczukii, S. kesselbrenneri, S. kurtzianum, S. lanciforme, S. lapazense, S. lechnoviczii, S. leptostygma, S. ligustrinum, S. longipedicellatum, S. luteum, S. macolae, S. macrocarpon, S. maglia, S. malinchense, S. mamilliferum, S. marginatum, S. mauritianum, S. melongena, S. miniatum, S. mochiquense, S. multidissectum, S. muricatum, S. neocardenasii, S. nigrum, S. nitidibaccatum, S. ochroleucum, S. okadae, S. oplocense, S. ottonis, S. pampasense, S. parodii, S. penelli, S. photeinocarpum, S. phureja, S. pinnatum, S. pinnatisectum, S. platense, S. platypterum, S. polyacanthos, S. polyadenium, S. prinophyllum, S. quitoense, S. radicans, S. raphanifolium, S. rostratum, S. rybinii S. salamanii, S. saltense, S. sambucinum, S. sanctae-rosae, S. sarrachoides, S. scabrum, S. schenkii, S. schickii, S. semidemissum, S. simplicifolium, S. sinaicum, S. sisymbrifolium, S. sodomaeum, S. soukupii, S. sparsipilum, S. spegazzinii, S. stenotomum, S. stoloniferum, S. suaveolens, S. subandigenum, S. sucrense, S. tarijense, S.tenuifilamentum , S. tlaxcalense, S. tomentosum, S. toralopanum, S. triflorum, S. tuberosum ssp. andigena, S. tuberosum ssp. tuberosum, S. tuberosum 'Aquila', S. tuberosum 'Xenia N', S. utile, S. vallis-mexicae, S. vernei, S. verrucosum, S. villosum, S. violaceimarmoratum, S. wittmackii, S. wittonense, S. xanti, S. yabari and S. zuccagnianum.
Note Oxalis tuberose (Oca), has been extensively tested in host range tests by Sullivan et al. (2007) and has been declared a non-host on this basis.
Host Plants and Other Plants AffectedTop of page
|Datura stramonium (jimsonweed)||Solanaceae||Other|
|Lycopersicon pimpinellifolium (currant tomato)||Solanaceae||Other|
|Solanum aviculare (kangaroo apple)||Solanaceae||Other|
|Solanum gilo (gilo)||Solanaceae||Other|
|Solanum lycopersicum (tomato)||Solanaceae||Main|
|Solanum marginatum (white-edged nightshade)||Solanaceae||Other|
|Solanum mauritianum (tobacco tree)||Solanaceae||Other|
|Solanum melongena (aubergine)||Solanaceae||Main|
|Solanum nigrum (black nightshade)||Solanaceae||Other|
|Solanum quitoense (naranjilla)||Solanaceae||Other|
|Solanum sarrachoides (green nightshade)||Other|
|Solanum tuberosum (potato)||Solanaceae||Main|
Growth StagesTop of page Pre-emergence, Seedling stage, Vegetative growing stage
SymptomsTop of page
Potato cyst nematodes, in common with other cyst nematodes, do not cause specific symptoms of infestation. Initially, crops will display patches of poor growth and plants in these patches may show chlorosis and wilting. When the tubers are harvested there will be a yield loss and tubers will be smaller. To be confident that these symptoms are caused by potato cyst nematodes and to give an indication of population density, soil samples must be taken or the females or cysts must be observed directly on the host roots. In heavily infested soils, plants have reduced root systems and often grow poorly due to nutrient deficiencies and to water stress. Plants may senesce prematurely as they are more susceptible to infection by fungi such as Verticillium spp. when heavily invaded by potato cyst nematodes.
Direct damage to roots and the yield of tubers
The infective second stage juveniles of both G. rostochiensis and G. pallida respond to environmental conditions when hatching. There is a short period of time for the second stage juvenile to locate a host root and begin the process of invasion, usually just behind the root tip. The juveniles then position themselves next to the stele within the root where, after a few hours, they will establish a feeding site (syncytium), which will become their nutrient source until their death. If a susceptible variety of potato is planted the plants will soon show signs of attack particularly when nematode density is high. In resistant plant varieties juveniles still hatch from the cyst and invade the plant roots, but they are unable successfully to establish a feeding site or syncytium. In this situation, males are more likely to be produced than females, as males have negligible nutrient requirements compared to females. Nevertheless, even resistant crops may show signs of attack.
The reduction in the yield of potato tubers, depending on the cultivar grown, is also related to or dependent on the plant's ability to tolerate the effects of nematode attack. The effects of potato cyst nematode on the plant include water stress and early senescence of the leaves. A heavily infested plant is unlikely to produce 100% ground cover with its reduced canopy of leaves. Many field studies have monitored the progression of ground cover by leaves and correlated the findings with yields (see Trudgill et al., 1998).
List of Symptoms/SignsTop of page
|Leaves / abnormal colours|
|Leaves / wilting|
|Roots / cysts on root surface|
|Roots / reduced root system|
|Vegetative organs / surface cracking|
|Whole plant / dwarfing|
|Whole plant / early senescence|
Biology and EcologyTop of page
Eggs contained within cysts are the persistent stage of the life cycle; new cysts contain around 500 eggs. Some eggs are able to survive within the cyst for as long as 30 years, although by this time very few are viable. Hatching in water is normal, especially within the 'hatching season' when most of the environmental factors are stable. Hatching is increased by the stimulus of a hatching factor, found in host root diffusate (Perry and Beane, 1988). This appears to affect the permeability of the eggshell lipoprotein membrane (Atkinson and Ballantyne, 1977a, b).
Many workers have studied the hydration of second-stage juveniles (Clarke and Perry, 1985; Perry, 1986; Perry and Feil, 1986; Holz et al., 1998) and pre-hatch behaviour within the egg (Doncaster and Shepherd, 1967; Doncaster and Seymour, 1973), as well as the responses of potato cyst nematodes to various hatching agents at different concentrations (Clarke and Hennessy, 1987). Studies have also been made of gene expression at hatching (Jones et al., 1997). Other factors affecting hatching are soil moisture, aeration and pH (Shepherd and Clarke, 1971). Soil temperature or the rate of heat accumulation (Jones and Parrot, 1969) are also important and Jones (1975) showed that the development of the nematode is governed by the rate of heat accumulation, except at extreme temperatures.
Nematode densities remain low at low temperatures (Mugniery, 1978; Franco, 1979; Hominick, 1979; Inagaki, 1984). The optimum temperature for the hatch of G. rostochiensis is about 15°C, with the largest proportion of adults in a population at 650-830 day degrees over a basal temperature of 4.4°C (Evans, 1968). Some populations seem to have adapted to climates with lower temperatures, as in the Scottish population from Ayrshire, which is able to mature even on early harvested potatoes (Ellenby and Smith, 1975; Hominick, 1979, 1982). The most northerly known point of survival for G. rostochiensis is the Finnish Arctic Circle after several years with long warm summers (Sarakoski, 1976).
The infective juveniles, as with most other cyst nematodes, penetrate the host root just behind the root tip. From this point the juveniles move up or down the root until they receive a specific signal, probably of a chemical nature, to set up a feeding site in the form of a syncytium. The ultrastructure of the feeding site and nematode interaction with the syncytium have been studied intensively (Wyss and Zunke, 1986; Golinowski et al., 1997; Endo, 1998). Infective second-stage juveniles that penetrate the pericycle cells of the plant are more likely to become males, whereas those that penetrate the procambial cells tend to become females (Golinowinski et al., 1997). Within a few hours of settling, the juvenile probes the selected cell, inserts its stylet into it while remaining motionless for several hours; the stylet is then withdrawn and re-inserted into the same cell. A secretory product from the oesophageal glands is injected via the stylet into the selected cell. The initial syncytial cell (ISC) is now altered to provide large amounts of nutrients to the developing nematode. The syncytium undergoes major changes, for example lysis of inner cell walls, formation of cell wall ingrowths next to plant conductive tissue, appearance of numerous lipid bodies and enlargement of now amoeboid nuclei. However, many events are still not fully understood. In resistant plants, the juvenile may try to form a syncytial feeding site but the walls of cells involved usually thicken and cells may die. This prevents the ready movement of nutrients to the juvenile (Rice et al., 1986; Robinson et al., 1988).
The life cycle takes approximately 45 days, during which time the males will moult and become vermiform, leave the host root and fertilize as many females as possible before dying about 10 days or so after first leaving the root (Evans, 1970). The females during this time have become saccate and their posterior ends have protruded through the root cortex, ready for mating.
Globodera spp. do not produce egg sacs or form a gelatinous mass into which eggs are deposited. However, it is known that pheromones are secreted which attract males (see data sheet on Globodera tabacum) (Green and Miller, 1969; Green and Plumb, 1970; Mugniery, 1979; Mugniery et al., 1992). Further studies into the nature of the pheromones (considered to be polar compounds by Green, 1980) have used electrophysiological techniques and behavioural assays (Riga et al., 1997). Extracellular recordings of electrical activity within the cephalic region of individual males were taken and correlated with exposure to extracts of females. The males exhibited a specific reaction to pheromones of females of G. rostochiensis, and it is also known that they will mate with females already fertilized. Trudgill (1967) noted that more males are present in a population when environmental conditions are poor, for example, lack of feeding sites due to overcrowding; the vermiform males are not known to feed (Evans, 1970).
Raising antibodies to nematodes permits investigations that were not previously possible (Fox and Atikinson, 1985b; Curtis, 1996), particularly studies on nematode secretions and their function and interaction within the host plant (Jones and Robertson, 1997; Jones and Harrower, 1998). The role of cuticular secretions in plant parasites is not yet fully understood (Forrest et al., 1989). Changes occur in the cuticle of second-stage juveniles when they begin to feed: the cuticle will alter due to natural growth patterns and, with the onset of moulting, changes occur that appear to be linked to interaction with the plant host (Jones and Robertson, 1997).
Recent observations show that one of the free radicals produced by plants is in the form of hydrogen peroxide, which is produced in response to nematode invasion and is in all probability broken down by an enzyme secreted by the nematode’s surface coat, thioredoxin peroxidase. Various other proteins have been identified from the surface coat and hypodermis of the nematode, such as the lipid binding protein GpFAR-1. This probably plays a role in the host defence signalling pathways as other plant parasitic nematodes, but not free-living types, also have similar compounds in their genetic makeup. Plant parasitic nematodes have probably evolved similar methods with which to protect and help to conceal themselves from the host plant.
Studies to explore the function of nematode genes, using the molecular tool RNAi, have given detailed insights into the host parasite relationship. Details of some of these findings regarding plant parasitic nematodes can be found in a review by Lilley et al., 2007. Genes related to the production of pectate lyases used for host penetration by G. rostochiensis (Kudla et al., 2007) provide an insight into the complexity of studies now being undertaken to elucidate aspects of the plant-host relationship.
Many studies are now focused on the interactions of chemical pathways, genetic function and metabolic concepts too numerous to cover here. Molecular aspects of plant-nematode interactions are reviewed by Gheysen and Jones (2006).
ClimateTop of page
|B - Dry (arid and semi-arid)||Preferred||< 860mm precipitation annually|
|BS - Steppe climate||Preferred||> 430mm and < 860mm annual precipitation|
|C - Temperate/Mesothermal climate||Preferred||Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C|
|Cf - Warm temperate climate, wet all year||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year|
|Cs - Warm temperate climate with dry summer||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers|
|Cw - Warm temperate climate with dry winter||Preferred||Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)|
|D - Continental/Microthermal climate||Tolerated||Continental/Microthermal climate (Average temp. of coldest month < 0°C, mean warmest month > 10°C)|
|Ds - Continental climate with dry summer||Tolerated||Continental climate with dry summer (Warm average temp. > 10°C, coldest month < 0°C, dry summers)|
|ET - Tundra climate||Tolerated||Tundra climate (Average temp. of warmest month < 10°C and > 0°C)|
Latitude/Altitude RangesTop of page
|Latitude North (°N)||Latitude South (°S)||Altitude Lower (m)||Altitude Upper (m)|
Air TemperatureTop of page
|Parameter||Lower limit||Upper limit|
|Absolute minimum temperature (ºC)||-80|
|Mean annual temperature (ºC)||10||25|
|Mean maximum temperature of hottest month (ºC)||27|
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
Means of Movement and DispersalTop of page
Generally, cyst nematodes build up in numbers over time, usually in patches. When the areas of infestation are disturbed i.e. by agricultural machinery, implements or farm workers, the cysts often adhering to clods of soil are easily transferred to uninfested areas. This can occur at all levels, locally and internationally. Transportation of crop plants and tubers also requires extremely high levels of hygiene to prevent movement. Packaging materials can also harbour cysts, which will readily transfer to a new favourable environment.
Natural Dispersal (Non-Biotic)
Wind can have an impact by lifting and spreading cysts in dry soil and plant debris from one area to another. Run-off flood water from infested areas can pose a threat as dry cysts float and can be carried some distance from the original source to start new infestations in local situations.
Vector Transmission (Biotic)
It is known that cyst nematodes are capable of passing through the digestive tract of farm animals and are excreted intact in a viable condition ready to begin a new infestation. Soil debris transported on animal hoofs can move cysts from one area to another.
Probably the most likely cause of new introductions to different countries has been unintentional and has come with the advent of trade and travel.
Pathway CausesTop of page
|Breeding and propagation||Potato breeding material from South America to Ireland||Yes||Yes||Turner and Evans, 1998|
|Crop production||Peru to Europe||Yes||Yes||Turner and Evans, 1998|
|Digestion and excretion||Yes|
|Flooding and other natural disasters||Yes||Been and Schomaker, 2006|
|Food||Yes||Yes||Turner and Evans, 1998|
|Garden waste disposal||Yes|
|Military movements||Yes||Yes||Brodie, 1998|
|People sharing resources||Yes|
|Seed trade||Netherlands to South America||Yes||DAO et al., 1971|
Pathway VectorsTop of page
|Clothing, footwear and possessions||Cysts||Yes||Been and Schomaker, 2006|
|Containers and packaging - non-wood||Cysts||Yes||Inagarki, 2004|
|Containers and packaging - wood||Cysts as contamination||Yes|
|Land vehicles||Cysts as contamination, juveniles||Yes||Yes|
|Machinery and equipment||Cysts||Yes||Yes||Been and Schomaker, 2006|
|Plants or parts of plants||Cysts, juveniles, eggs||Yes||Yes|
|Soil, sand and gravel||Cysts in soil and dust storms, juveniles||Yes||Yes|
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bulbs/Tubers/Corms/Rhizomes||cysts||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Growing medium accompanying plants||cysts||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Roots||cysts; eggs; juveniles||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Seedlings/Micropropagated plants||cysts; eggs; juveniles||Yes||Yes||Pest or symptoms not visible to the naked eye but usually visible under light microscope|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|True seeds (inc. grain)|
Impact SummaryTop of page
Economic ImpactTop of page
The actual cost of damage caused by G. rostochiensis is difficult to determine, but potato cyst nematodes (see also Globodera pallida) cause extensive damage, particularly in temperate areas and particularly when virulent pathotypes occur and any resistance has failed. The situation is worse with G. pallida, where commercial cultivars with good resistance are few and often have other undesirable properties. Damage is related to the number of eggs per unit of soil, and is reflected in the weight of tubers produced (see Seinhorst, 1986 for further details). Severe infestation with G. rostochiensis and/or G. pallida can result in yields that are smaller than the quantity of seed originally planted (Oerke et al., 1994).
South and Central America
In Chile, microplot studies were carried out to determine the effect of densities of G. rostochiensis (as eggs/g soil) on the yield of summer, winter and spring sown potatoes (Moreno et al., 1984; Greco and Moreno, 1992). For the summer crop, a damage threshold for tuber yield of 1.3 eggs/g soil was determined. At population densities greater than this, yields were greatly suppressed. At initial inoculum levels of 12, 32 and 128 eggs/g soil, reductions in yield of 20, 50 and 70%, respectively, were recorded. Winter-sown potatoes indicated a tolerance limit of 0.8 eggs/g soil. Spring-sown potatoes had a tolerance limit of 1.56 eggs/g soil. Yield losses of 20, 50 and 90% were obtained with population densities of 9, 28 and 128 eggs/g soil, respectively.
In Panama, Pinochet (1987) estimated that average annual crop losses due to G. rostochiensis, G. pallida and species of Meloidogyne were 10-30%.
In Canada, G. rostochiensis was found in Newfoundland in 1962, followed by G. pallida 15 years later. Since their discovery, ca $Can 800,000/year has been spent on control and research (Miller, 1986). In 2006, G. rostochiensis was discovered in a field in the Saint-Amable region of Quebec, Canada. (Sun et al., 2007).
Between 1967 and 1973 in the UK, Brown and Sykes (1983) investigated the losses due to G. rostochiensis and G. pallida in Staffordshire and Worcestershire. At densities of 0-24 eggs/g soil, losses of 6.25 t/ha per 20 eggs/g soil were recorded. At 40-160 eggs/g soil, losses of 1.67 t/ha were recorded. An average loss of 2.75 t/ha per 20 eggs/g soil was determined for all population densities. The maximum crop loss was 22 t/ha.
The economic threshold for G. rostochiensis in the UK is ca 15 eggs/g soil. Unless control measures are implemented above this threshold, crop losses can be as high as 80%. Dale (1988) found that, on average, the losses are 3-4 t/ha per 10 eggs/g soil at planting time. A loss figure of 9% nationwide does not take into account any indirect losses sustained as a result of control or quarantine measures, or for growing crops which give a lower return (Evans and Brodie, 1980; Brodie, 1984). The annual UK expenditure on nematicides is £9 million and these are applied over ca 40,000 ha, a cost of £210-250/ha; losses in spite of control measures are ca £6 million (Trudgill et al., 1987; Dale, 1988).
In Germany, Engel et al. (1982) estimated the marketable crop losses dependent on G. rostochiensis infestation density in long-term trials using regression analysis. They estimated crop losses of 11, 27 and 43% at nematode populations of 100, 1000 and 10,000 larvae/100 cm³, based on an attainable yield of 22.4 t/ha on nematode-free plots. As populations of 500 juveniles/100 cm³ are regularly reached, losses on affected areas were estimated at 10-15% (Kleinhempel, 1986).
Greco et al. (1982) reported that the loss threshold for G. rostochiensis and G. pallida at three Italian sites was between 1.4 and 2.1 eggs/g soil. In a field trial in 1982, a yield of 23.9 t/ha was achieved on nematicide treated areas. Without control, G. rostochiensis reduced the yield by 76%; the marketable yield of 13.75 t/ha on treated areas was reduced (by 85%) to 1.96 t/ha (Greco et al., 1984).
In Arcadia, Greece, application of nematicides to control G. rostochiensis increased the yield of potatoes grown on sandy soils to 31 t/ha. In untreated areas, the loss was 37% (Kalyviotis-Gazelas, 1982).
In Poland, monoculture has been used as a method of growing potatoes. This has led to a build-up in the population of nematodes, particularly G. rostochiensis. Zawislak et al. (1981) reported that after growing potatoes in the same ground in Balayny for 4 years, the number of viable cysts rose to 63/100 g soil, though yield was unaffected. In the fifth year, however, there were 180 cysts/100 g soil (up to 3000 cysts/plant) and yields fell by 72%.
G. rostochiensis is a principal potato pest in Byelorussia. At a population of 1000 juveniles/100 cm³ soil, crop losses of 17-20% were recorded in susceptible varieties. At population densities of 500, 10,000 and 25,000 juveniles/100 cm³ soil, losses of 31, 45 and 74%, respectively, were recorded. With repeated cropping (as in the Polish monoculture systems), the population of G. rostochiensis could reach 38,000/100 cm³ (Gladkaja and Korzhentsevskaya, 1985).
Risk and Impact FactorsTop of page Invasiveness
- Invasive in its native range
- Proved invasive outside its native range
- Has a broad native range
- Abundant in its native range
- Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
- Pioneering in disturbed areas
- Highly mobile locally
- Long lived
- Fast growing
- Has high reproductive potential
- Has propagules that can remain viable for more than one year
- Has high genetic variability
- Changed gene pool/ selective loss of genotypes
- Host damage
- Increases vulnerability to invasions
- Modification of nutrient regime
- Modification of successional patterns
- Negatively impacts agriculture
- Negatively impacts cultural/traditional practices
- Negatively impacts livelihoods
- Competition - monopolizing resources
- Pest and disease transmission
- Interaction with other invasive species
- Parasitism (incl. parasitoid)
- Highly likely to be transported internationally accidentally
- Difficult to identify/detect as a commodity contaminant
- Difficult to identify/detect in the field
- Difficult/costly to control
Uses ListTop of page
- Laboratory use
- Research model
- Test organisms (for pests and diseases)
DiagnosisTop of page
Following the demonstration that isoelectric focusing could be used to identify G. rostochiensis and G. pallida because they display different protein profiles (Fleming and Marks, 1983), DNA based techniques of identification have been developed (Bakker and Gommers, 1986; Marshall and Crawford, 1987; Bossis and Mugniery, 1993; Thiery and Mugniery, 1996). More recently, these techniques have been improved upon and identification and the use of species-specific primers (Mulholland et al, 1996; Fullaondo et al, 1999), for RAPD PCR RFLP, and AFLP multiplex PCR are now recommended techniques for molecular identification. Micro array techniques are now also more commonly used for diagnostic work (Picard et al., 2007).
Molecular diagnostics are now the “norm” having advanced dramatically in the last decade. Their discriminatory powers can provide a deeper level of species separation and identification. Molecular diagnostics, although still comparatively expensive, require less skill then traditional methods and can give a greater throughput and standardisation of samples on a routine basis. It is also possible to determine the identity of known pathotypes of potato cyst nematode using a high performance capillary electrophoresis (CE) technique. This uses sodium dodecyl sulphate (SDS) capillary gel electrophoresis to display polypeptide profiles (Hinch et al., 1998). Other means of discriminating nematode species using biochemical techniques have led to the use of monoclonal antibodies as a diagnostic tool (Fox and Atkinson, 1985a; Curtis, 1996). Perry and Jones (1998) reviewed further in-depth biochemical and genetic studies, including the disruption of the nematode life cycle by the addition of resistance genes or the insertion of other genetic constructs into the plant genome by genetic engineering techniques (Burrows, 1996; Burrows and De Waele, 1997).
Detection and InspectionTop of page
Potato cyst nematodes, in common with other cyst nematodes, do not cause specific symptoms of infestation. Initially, crops will display patches of poor growth and these plants may show chlorosis and wilting. When the tubers are harvested there will be a yield loss and tubers will be smaller. To be confident that these symptoms are caused by potato cyst nematodes and to give an indication of population density, soil samples must be taken or the females or cysts must be observed directly on the host roots. Detection based on host plant symptoms and identification by morphological and molecular methods are detailed in EPPO (2009).
Surveys of the numbers and distribution of potato cyst nematode are prerequisites for making informed choices for their management. Samples taken within a field are either to check whether potato cyst nematode is present or not in the field for statutory purposes or to determine the extent of the infestation, which might include a determination as to what species is present.
At one time, it was considered that nematodes had a haphazard distribution in the field but this has been disproved. Aspects of the environment and ecological factors such as disease, predators and soil type favour aggregated distributions. Many models help to describe distributions, for example Taylor's Power Law (Taylor, 1961), Iwao's regression model (Allsopp, 1990) and others. However, geostatistical techniques may provide a more purposeful definition of the spatial distribution of nematodes. Although these techniques are young, 3-D maps can be generated to study nematode population levels more effectively. These methods have already proved useful in mapping other types of field related data (Chellemi et al., 1988) and have recently been applied to the distribution of potato cyst nematodes (Evans et al., 2003).
Similarities to Other Species/ConditionsTop of page
G. rostochiensis is very similar to the other members of the round cyst nematode group. Its morphology, phenotypic appearance and biology as well as host range can make difficult the initial diagnosis of the species from Globoderatabacum tabacum (Lownsbery and Lownsbery, 1954), Globoderatabacum solanacearum (Miller and Gray, 1972), Globoderatabacum virginiae (Miller and Gray, 1968) and Globoderapallida (Stone, 1972). G. rostochiensis, the G. tabacum 'complex' and Globodera spp. from Compositae all produce females that have a golden phase upon maturity just before death, after which the cyst cuticle of all becomes brown. The second-stage juveniles are similar in, for example, body and stylet length. Other studies have shown hybridisation of G. rostochiensis with the G. tabacum group, proving a strong genetic link. Their host ranges are also very similar (Roberts and Stone, 1981; Stelter, 1987). G. pallida is often found together with G. rostochiensis but has white or cream females, the second-stage juveniles are longer and have longer stylets and the cyst has a larger Granek's ratio and usually fewer ridges between the anus and fenestra. The two species can be differentiated using simple isoelectric focusing techniques. For details, see the Diagnosis section.
Prevention and ControlTop of page
To prevent further spread of potato cyst nematodes into uninfested areas, several methods are used. These include legislation on the movement of seed potatoes, nursery stock, flower bulbs and soil. These apply internationally and nationally.
The specific EPPO quarantine regulations (OEPP/EPPO, 1990; EPPO, 2007) for these nematodes require that fields in which seed potatoes or rooted plants for export are grown are inspected by taking soil samples according to an EPPO-recommended method (OEPP/EPPO, 1991; EPPO, 2007) and must be found free from viable cysts of both species of potato cyst nematode. The sampling must be performed after harvest and after removal of the previous potato crop.
Quarantine is a very necessary and often expensive way of attempting to limit the damage caused by disease organisms such as nematodes. Methods to limit or prevent the introduction of alien or existing pests by providing ways of very accurate identification plus sensible legislation (MAFF, 2000) are already used in many countries. Continuous records must be kept of previous land usage and crop rotations. Various organizations throughout the world help to back up the phytosanitary regulations.
The previous PCN Control Directive for the European Union was in place for several decades, over which time many practices within the potato industry have changed and a lot more has been learned from intensive studies on this pest. New European legislation was introduced on 1 July 2010. The revised Control Directive (2007/33/EC) was adopted by the Council of Ministers and has been published in the Official Journal of the European Communities in June 2007.
Physical barriers help to confine pests to their own locality, for example, seas, mountains and other natural phenomena. Trade is probably the keystone to the problem of spread (Parker, 2000). Trade is essential and, when it comes to the movement of soil and plants, is of unparalleled importance with regard to nematode quarantine. Plant parasitic nematodes occur worldwide on virtually every crop, and plant movement can probably be blamed for all potato cyst nematode occurrences outside South America.
1. Check that machinery is thoroughly clean and free from plant debris.
2. Do not return soil to fields as it may cause infestation of potato cyst nematode to spread.
3. Clean soil from potato tubers and have the soil tested to be sure of non-transference of potato cyst nematode.
4. Make sure that laboratories that test soil for potato cyst nematode are properly qualified and that they test 500 g of soil per sample.
5. Grow susceptible and resistant varieties of potato alternately, thus reducing the possibility of selecting a highly virulent or new pathotypes.
For further details, see Seinhorst (1986).
Rotation is frequently used to reduce population densities of the potato cyst nematodes, G. rostochiensis and G. pallida. The major hosts of these two species are restricted to the plant family Solanaceae, with the main commercial crops being potato, tomato and aubergine. When these crops are grown in monoculture for several seasons in infested soil, nematode densities can increase to extremely high levels and crop yields become uneconomical. To reduce nematode population densities, non-host crops such as barley are grown between host crops. Magnusson (1987) recorded an 87% decline in G. rostochiensis population density using this type of rotation. Whitehead (1995) also found good control of G. rostochiensis when barley was grown in infested microplots. The annual decline rate of potato cyst nematode in soil is variable, depending on the non-hosts used, the initial population density, various soil-related factors and the population under study. If the reduction of population densities by rotation alone is too slow, then additional means of control may be necessary, such as the use of resistant cultivars, trap cropping or nematicides.
Trap cropping has been used successfully for the reduction of cyst nematode populations (Halford, et al., 1999). Potatoes are grown in order to cause the second stage juveniles to hatch. These are given sufficient time to penetrate the roots and develop into young adults. By monitoring soil temperature from the date of planting, fertilization and formation of new eggs can be avoided by destroying the crop some 6 or even 7 weeks after planting, before too many heat units have accumulated. If crop destruction is left too late, the nematode density will increase. Using this method, G. rostochiensis populations have been reduced by more than 80% (Halford et al., 1999).
The use of resistant potato clones as trap crops has been used in field trials in Northern Ireland. Ten clones had a strong hatch-inducing effect and resistance to currently known PCN pathotypes. Some of these showed potential for further development to reduce high population levels of PCN in the field and for the organic potato market (Turner et al., 2006) Another solanaceous host, S. sisymbriifolium, acts in a similar way, stimulating hatch without the nematode being able to complete its life-cycle, with the added benefit of then being used as a green manure. The seed line of S. sisymbriifolium is probably important as seed lines used for trap cropping are listed by Scholtze (2000) as totally resistant to PCN, whereas two lines (nos. 72 and 121) were recorded as poor hosts (with less then 5 cysts per plant in a host test performed by Stelter (1987). Where the nematode density is reduced, there will be a significant yield benefit for any subsequent potato crop.
Solarization is a good method of killing nematodes in very hot climates. The soil is covered with two layers of polyethylene, allowing the soil underneath to heat up quickly. In Oman, Mani et al. (1993) found that 62 days of solarization reduced G. rostochiensis population density by 95%. In New York State, USA, 97% of nematode eggs were inactivated in the top 10 cm of soil (LaMondia and Brodie, 1984.). Solarization in cooler climates and at depths greater than 10 cm is much less effective.
Natural parasites and biological control options are being studied intensively in the search for natural ways of controlling plant parasitic nematode populations without the use of nematicides, which are highly toxic and a burden on the environment. These biological control agents are part of a grander objective to manage nematodes more effectively using a variety of biological strategies that include trap cropping and rotation.
Work on biological control agents was started in the late 1930s (Linford et al., 1938) and still continues (Crump and Flynn, 1995; Segers et al., 1996; Crump, 2004). At present, there is still no commercial biological product available to control potato cyst nematodes. The majority of studies in the late 1990s have concentrated on the fungal control agents Pochonia, Hirsutella and Arthrobotrys and the bacterium Pasteuria.
Several workers (e.g. Roessner, 1986) have studied biological control of G. rostochiensis in pots and in vitro. Almost no field trial data are available. This is probably due in part to the logistics of such operations e.g. producing enough inoculum for field scale trials. Also, some tests do not produce the expected results for reasons as yet undefined, but are probably related in some way to the physiology or ecology of the nematode or to the host parasite relationship.
Pochoniachlamydosporia will infect young females in pots but is less effective when potatoes are grown in the presence of low nematode population densities.
Progress in the area of biological control requires a better understanding of the population dynamics of potato cyst nematodes and its parasites (Davies et al., 1991; Davies, 1998). A variety of factors, such as plant host, the action of root exudates, soil type and the mode of parasitism of the control organism, interact to determine success in the biological control of the nematode. Cyst nematodes may be more susceptible to infection at certain points in their life cycle. For example, the three major fungal parasites Pochonia chlamydosporia, Fusarium oxysporum and Cylindrocarpon destructans, have all been detected throughout the potato cyst nematode life cycle but the most active will vary at different times of the cycle (Crump, 1987).
At the molecular level, the high specificity known to occur between cyst nematodes and their plant hosts is important, in terms of intra- and inter-specific variation, particularly with regard to virulence and avirulence. Work by Segers et al. (1996) on the effect of a protease-like enzyme (designated VCPI), from the nematophagous fungus Pochonia chlamydosporia, showed that when this enzyme was used as a pre-treatment of Meloidogyne incognita eggs they became more susceptible to P. chlamydosporia. The same treatment applied to G. rostochiensis eggs gave no response.
In the last decade some products have come to market that have nematicidal effects, such as DiTera, a compound produced from the fermentation extracts of a bacterium. Paecilomyces chlamydosporia, a fungal biocontrol agent, is also available on the market. Most other potential biocontrol agents are still being tested or studied to overcome problems with delivery systems or application methods. Many other mutualistic bacterial and fungal endophytes probably exist in the agroecosystem that would greatly improve plant health while at the same time be detrimental to plant parasitic nematodes. However, many technologies are involved in discovering the most appropriate candidates for commercialisation (Sikora et al., 2007). With time, appropriate study of plant parasites, their molecular properties and modes of parasitism will improve biological control options and identify new ways forward.
Fumigant nematicides are toxic and expensive, but have been used to help reduce large densities of potato cyst nematode. Soil fumigants can kill large numbers of nematodes, especially in moist sandy soils under polythene sheeting. Soil fumigants can be injected into the soil and this is also very effective, but there may be some risks of ground water contamination in certain circumstances. Many chemicals previously used for nematode control have now been banned, or are in the process of being phased out, due to safety concerns or their effect on the ozone layer.
The nematicide 1, 3-Dichloropropene is available mixed in various quantities with other compounds, such as C3-chlorinated hydrocarbons and 1, 2-dichloropropane; Telone II consists of 94% 1, 3-D whereas D-D contains around 50%. Other types of fumigants release methyl isothiocyanate (MITC), e.g. dazomet, which is effective against potato cyst nematode (Whitehead, 1975).
Fumigant nematicides are usually applied several weeks before planting to avoid phytotoxicity. Conditions affecting the efficacy of any of this type of chemical include soil type, moisture content, drainage and temperature.
For further information, see Whitehead (1998), Whitehead and Turner (1998) and Haydock et al. (2006).
These compounds are used in smaller amounts and usually do not persist in the soil as long as fumigant nematicides. Organophosphates and oximecarbamates are very effective nematicides. Their effect on nematodes is to paralyse rather than kill, unless very large doses are used. Organophoshates offer good control of G. rostochiensis, but need to be incorporated into the soil using rotary cultivation. These types of chemical are better suited to light, silty soils and are not as effective on organic soils. Isazophos (CGA 12223) was used by Moss et al. (1975) on silty loams and gave partial control of G. rostochiensis and of a mixed population of G. rostochiensis and G. pallida.
Organophosphates are cumulative poisons and, as such, are not used as much as they once were. The pH of the soil is an important factor if used: at more than pH 8.0 the soil is too alkaline. Higher temperatures also cause degradation of carbamates at faster rates than at lower temperatures.
Non-fumigant chemicals are most effective when used in granular form and at 15 cm. below the soil surface incorporated by machines that won't damage the soil structure.
Ellenby (1954) was the first to report a gene for resistance to G. rostochiensis. The gene was found in Solanum tuberosum ssp. andigena and is a single dominant gene, referred to as gene H1, but it conveys resistance only to certain populations of G. rostochiensis. Some European cultivars of potato have resistance (often only partial) to European pathotypes of potato cyst nematode but some South American populations of potato cyst nematode are known to be more virulent and are able to override resistance in the European cultivars (Kort and Jaspers, 1973; Turner et al., 1995).
Intensive studies in South America, at the International Potato Centre (CIP) in Lima, Peru, over the past 20 years, have considered 3000 accessions of potato and have focused on pathotypes in the local regions. Some resistant cultivars reduce potato cyst nematode field populations by up to 50%, but may select new, more virulent pathotypes. The use of any resistant potato cultivar repeatedly carries the risk of selecting a virulent nematode population, perhaps even a type not previously recognized, especially where unknown mixtures of species and/or pathotypes occur in the same location. It is wise to grow a different cultivar (resistant or tolerant) each year to avoid this problem. In the UK, frequent growing of the cultivar Maris Piper, resistant to UK G. rostochiensis potato cyst nematode populations but susceptible to G. pallida, has caused UK populations to switch to G. pallida, which is much more difficult to control.
In Asia both G. pallida and G. rostochiensis are present in India, Pakistan and Sri Lanka but growing areas are restricted; so far only the R1A and P5A pathotypes are known to be present (Zaheer, 1998). In South Africa, the potato is a major crop with some 5600 ha grown, most of which are sold for consumption. In Morocco, 5 eggs/ml soil caused appreciable damage (Spears, 1968).
There is also a need for tolerant cultivars, which suffer less damage and growth of which may also prevent selection of virulent pathotypes by resistant cultivars in a field. Strict quarantine measures are important supporting measures. There are no potato cultivars with full resistance to G. pallida (Marshall, 1998). However, in New Zealand the Institute for Crop and Food Research has released a series of cultivars with high resistance to G. rostochiensis and G. pallida, e.g. Karaka (Anderson et al., 1993) and Gladiator (Genet et al., 1995). For other resistant, tolerant and partially resistant cultivars, see Whitehead (1998).
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18/03/2008 Updated by:
Janet Rowe, IACR-Rothamsted, Rothamsted Experimental Station, Harpenden, Hertfordshire, AL5 2JQ, UK
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