What Are Cereal Cyst Nematodes – How To Stop Cereal Cyst Nematodes

By: Becca Badgett, Co-author of How to Grow an EMERGENCY Garden

Most wheat,oatsand barleyvarieties grow during cool seasons and mature as the weather warms. Growingfrom early winter with a late spring harvest, the crop is less vulnerable towarm season pests. However, there are issues that arise during the cool season.One of the most prominent issues is cereal cyst nematodes. If you’re curious andasking, “what are cereal cyst nematodes,” read on for an explanation.

Cereal Cyst Nematode Info

Nematodes are tiny worms, often roundworms and cutworms.Some are free-living, feeding on plant materials such as wheat, oats andbarley. These can cause extreme damage and make crops unsaleable.

Yellowing patches above ground can indicate you have thisnematode in the crop. Roots may be swollen, ropey or knotted with shallowgrowth. Small white cysts on the root system are female nematodes, loaded withhundreds of eggs. Juveniles do the damage. They hatch when temperatures dropand autumn rain occurs.

Warm and dry weather in fall delay hatching. These nematodesdo not usually appear and develop until after the second planting of a cerealcrop in the same field.

Cereal Cyst Nematode Control

Learn how to stop cereal cyst nematodes to avoid such issueswith your crops. A few ways to do this include:

  • Plant early to allow a good root system to develop.
  • Grow resistant types of cereal cultivars to limit chances of nematodes.
  • Rotate crops every year or two. The first planting seasons are not usually when cereal cyst nematodes occur. If a serious infestation occurs, wait two years before planting a cereal crop in the spot again.
  • Practice good sanitation, keeping weeds out of your rows as much as possible. If you plant an alternate crop in the same spot in summer, keep weeds down then as well.
  • Amend soil to improve drainage and keep the soil as fertile as possible.

Fertile, weed-free and well-draining soil is less likely toretain these pests. Cereal cyst nematodes only feed on grasses and cereal cropsand use those plants for hosts. Plant a non-cereal crop in spring to encouragethose remaining to move out because of no host and food shortage.

Once your field is infested, cereal cyst nematode control isnot practical. It is highly dangerous to use chemicals on these crops and thecost is prohibitive. Use the tips above to keep your field free of the pest.

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Host induced gene silencing for nematode resistance in plants. ( A ) and ( B ): Double stranded RNA (dsRNA) sequence of the target nematode effector gene is transformed into the plant which is cut by the plant DICER enzyme into small interfering RNAs (siRNAs). ( C ), ( D ) and ( E ): These siRNAs are taken up by the nematodes through their stylets which are detected by the RISC complex that binds to mRNA (complementary to the siRNA sequences) of the target effector gene. ( F ) and ( G ): This is followed by the activation of nematode DICER to cut the double stranded RNA to degrade the mRNA of the target nematode effector gene. Reproduced from Ali et al. [ 10 ].

Heterodera ciceri (chickpea cyst nematode)


Heterodera ciceri (chickpea cyst nematode)


  • Pictures
  • Identity
  • Taxonomic Tree
  • Notes on Taxonomy and Nomenclature
  • Description
  • Distribution
  • Distribution Table
  • Risk of Introduction
  • Habitat
  • Hosts/Species Affected
  • Host Plants and Other Plants Affected
  • Growth Stages
  • Symptoms
  • List of Symptoms/Signs
  • Biology and Ecology
  • Natural enemies
  • Pathway Vectors
  • Plant Trade
  • Impact
  • Detection and Inspection
  • Similarities to Other Species/Conditions
  • Prevention and Control
  • References
  • Distribution Maps


  • Last modified
  • 10 December 2020
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Preferred Scientific Name
  • Heterodera ciceri
  • Preferred Common Name
  • chickpea cyst nematode
  • Taxonomic Tree
  • Domain: Eukaryota
  • Kingdom: Metazoa
  • Phylum: Nematoda
  • Class: Secernentea
  • Order: Tylenchida

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TitleSEM of H. ciceri
CaptionFemale neck and excretory pore (arrowed). Scale bar = 100µm.
CopyrightFranco Lamberti/CNR - Istituto di Nematologia Agraria
TitleSEM of H. ciceri
CaptionTerminal cone, a=anus. Scale bar = 100µm.
CopyrightFranco Lamberti/CNR - Istituto di Nematologia Agraria
TitleSEM of H. ciceri
CaptionUnfenestrated vulval cones with interlacing cuticular pattern extending over large distance into vulval areas. Scale bar = 10 µm.
CopyrightFranco Lamberti/CNR - Istituto di Nematologia Agraria
TitleSection of chickpea root showing infestation
CaptionRoot infested with H. ciceri and showing the nematode females (n) and syncityal cells (s) occupying nearly the entire stelar portion and pressing on xylem (xe) and phloem elements.
CopyrightNicola Greco
TitleInfested chickpea root
CaptionChickpea root heavily infested with H. ciceri. Note the several females (f) and cysts (c) of the nematode, surrounded by small necrotic areas (A) and the lemon shape of these (B). Scale bar = 450 µm.
CopyrightFranco Lamberti/CNR - Istituto di Nematologia Agraria
TitleInfested lentil root
CaptionLentil root showing numerous females (f) of H. ciceri. Note the absence of the necrotic areas on the root around the females.
CopyrightNicola Greco
TitleInfested chickpea roots
CaptionChickpea roots infested with H. ciceri. A) Superficial necrotic areas (arrowed) at the penetration loci of second stage juveniles. B) Necrotic symptoms and a fourth stage female (arrowed) rupturing the rhizoderm.
CopyrightNicola Greco
TitleDamage symptoms on chickpea crop
CaptionA chickpea crop in north Syria showing extensive yellowing caused by severe infestation of H. ciceri.
CopyrightNicola Greco
TitleDamage symptoms on chickpea crop
CaptionEffects of increasing (right to left) soil population densities of H. ciceri on the growth of winter chickpea (foreground) and lentil (background). Note that the severity of the same nematode density is much more on chickpea than on lentil. Numbers are eggs of the nematode/cm³ soil.
CopyrightNicola Greco


Preferred Scientific Name

Preferred Common Name

EPPO code

Taxonomic Tree

  • Domain: Eukaryota
  • Kingdom: Metazoa
  • Phylum: Nematoda
  • Class: Secernentea
  • Order: Tylenchida
  • Family: Heteroderidae
  • Genus: Heterodera
  • Species: Heterodera ciceri

Notes on Taxonomy and Nomenclature


As with all species of the genus, H. ciceri has four juvenile stages, the adult stages and finally the cyst stages. Cysts contain many eggs (100-400). Sexual dimorphism is very marked.

Length 134 (123-143) µm width 50 (48-53) µm length/width ratio 2.5 (2.4-2.8) egg shell unsculptured and hyaline. Second stage juvenile folded four times within the egg shell.

Vermiform. Length 525 (440-585) µm stylet length 28.6 (27-30) µm maximum width at mid body 21 (19-22) µm tail width at anus 14-15 µm, tail length 60 (53-72) µm length of hyaline tail portion 36 (31-42) µm head tip to median bulb distance 82 (80-86) µm head tip to excretory pore distance 118 (105-128) µm lateral field width 4-5 µm, 1/4 of body width dorsal gland orifice 5-6 µm behind stylet knobs. Head hemispherical slightly offset an oral disc plate dorso-ventrally elongated and two rounded lateral sectors bearing large semilunar amphidial apertures. Stylet robust, knobs 2-2.5 µm long, 4-5 µm wide with concave anterior surfaces. Oesophageal glands well developed, 37% (30-42) body length from head. Hemizonid distinct and two annules long, 1-2 annules anterior to excretory pore. Oval genital primordium, 13-15 µm long, 10-13 µm wide, 218 (210-232) µm anterior to tail tip. Cuticular annulations distinct. Lateral field with four incisures, 20-25% of body width having outer bands aerolated. Tail irregularly annulated, tapering uniformly and rather abruptly to a finely rounded terminus, with a terminal hyaline portion 54 (48-58)% of tail length.

Length, excluding neck, 773 (550-950) µm width 451 (300-520) µm length/width ratio 1.7 (1.5-2) stylet length 30 (29-31) µm dorsal gland orifice 5-6 µm excretory pore 158 (154-169) µm from anterior end. Body typically lemon-shaped with well defined neck and prominent terminal cone. Adult female opaque-white in colour, turning yellow and then yellowish-brown at the start of the tanning. Occasionally a thin sub-crystalline layer may cover the entire female body. Gelatinous matrix present after the females becomes gravid, but only very seldom containing 1-2 eggs. Vulva a terminally positioned transverse slit. Anus subterminal, located in a depression 7-10 µm in diameter. The cuticle is ornamented by zigzag external ridges at mid body, but ornamentation of the neck region and in the cone consists of a series of unbroken lines which, in scanning electron microscopy end-on view, appears as a number of concentric circles surrounding the vulva area.

Length, excluding neck, 796 (570-930) µm width 452 (350-550) µm ratio 1.77 (1.56-2.37) neck 115 (90-160) µm. Cysts are typically lemon-shaped with a distinct neck and a prominent cone. Neck may be offset and curved posteriorly. Cysts well ornamented with irregular zigzag ridges, forming a fine network over the body. A radial pattern of 15-20 straight ridges on the apex of the vulval cone, including the semifenestral areas before fenestration. Excretory pore 10-14 µm across within a depression and encircled by 3-4 continuous cuticular ridges. Cysts covered partially or entirely with a thin, white subcrystalline layer.

Cysts ambifenestrate semifenestrae semicircular and sub-equal.

Fenestral length 40 (32-52) µm maximum fenestral width 27 (20-37) µm semifenestral length 17 (13-20) µm length of vulval slit 50 (43-60) µm. Numerous prominent, dark-brown bullae irregularly distributed at the periphery of the vulval cone at and below the underbridge level in most cysts. Underbridge well developed 125 (115-160) µm long, lying 78-86 µm below apex of the vulval cone, with furcate ends and central thickening.

Length 1308 (1235-1448) µm maximum body width 29-30 µm stylet length 29 (28-30) µm spicules length along axis 36 (34-38) µm gubernaculum 9-10 µm head tip to excretory pore distance 170 (153-180) µm. Males are abundant in this species. They are vermiform, typically an open-C-shaped after heat relaxation, with a short, bluntly rounded tail, about 1/3 of body width length, and with a long (10-14 µm) cloacal tube ending in a circular opening. Head region hemispherical, 5-6 µm long and 10 µm wide, offset from the body. Prestoma rectangular shaped, situated in the centre of the slightly raised oral disc plate. Labial disc plate partially fused with the rounded subdorsal and subventral pairs of lip sectors. Large amphidial apertures lie between the labial disc and the ellipsoidal lateral sectors. Posterior to the lip sectors the head annules are irregular and incomplete, except the basal annule which is large and less subdivided by longitudinal striae. Cephalic framework robust stylet also robust with rounded basal knobs 5-6 µm wide. Dorsal gland orifice opens 5-6 µm behind stylet knobs. Median oesophageal bulb oval, 25 x 10 µm, with moderately developed valve 98-117 µm from head tip. Hemizonid conspicuous, 2 annules long, located 9-12 annules anterior (13-21 µm) to the excretory pore. The excretory pore is located 170 (153-180) µm from the anterior end and is visible by scanning electron microscopy as a small pore about half an annule width in diameter. Testis single not reflexed and occupies 61-68% of the body length. Spicules arcuate, tapering distally. Gubernaculum with slight ventral curve, not ornamented. The lateral field has four equidistant incisures and is about 1/4 of the body width at mid-body. Anteriorly the field begins at the 6-8th body annules as three lines forming two regularly areolated bands.


Distribution Table

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.


Risk of Introduction


Hosts/Species Affected

Host Plants and Other Plants Affected

Growth Stages


The major symptoms on the roots of chickpea are the presence of white and yellow females at the flowering to early podding stage and of brown cysts at later stages (late podding to senescence). At earlier plant stages and in the case of heavy infestations, small necrotic spots (2-3 mm long and 0.5 mm wide), caused by juvenile penetration, appear longitudinally on the roots, from the centre of which the fourth and adult stage of the nematode will protrude. Rhizobium nodulation of heavily infested roots is reduced.

Symptoms on aerial plant parts are not specific they are those characteristic of a plant whose root system is damaged so that water and nutrient uptake is impaired. Plants with severely infested roots are stunted, yellowish, senesce earlier and have limited number of flowers and pods, which may be empty. The severity of these symptoms is correlated with intensity of the population density of the nematode in the soil and generally infestations become obvious from early flowering onwards.

List of Symptoms/Signs

SignLife StagesType
Growing point / dwarfing stunting
Leaves / yellowed or dead
Roots / galls along length
Roots / necrotic streaks or lesions
Stems / discoloration
Whole plant / discoloration
Whole plant / dwarfing
Whole plant / early senescence

Biology and Ecology

The nematode survives in the soil as a cyst in the absence of host plants or during unsuitable environmental conditions. It normally develops from winter to spring, but on the Anatolian Plateau in Turkey it also develops through the summer. When a host plant is sown, usually from late autumn to early spring, the nematode is stimulated to hatch by root exudates. Second stage juveniles, emerging from eggs contained within cysts, are the only infective stage they migrate in the soil and when a root is encountered they penetrate near the tip and move within it for a few mm. They then insert their heads in the vascular cylinder, become sedentary and stimulate the formation of syncytial cells around their cephalic region on which they feed. The nematode then undergoes three moults becoming third and fourth stage juveniles and then adults. The life span of the different stages is greatly influenced by environmental conditions. After fertilisation, the female produces a small egg sac, made of a gelatinous matrix, and starts to produce eggs. All eggs are retained within the female body and very occasionally a few (1-2) are laid in the egg sac. When females are old or environmental conditions become unsuitable (temperature around 30°C, low soil moisture content, host plant senescing), the female turns into a cyst by thickening its cuticle, which also becomes brown, and thus protects the eggs contained inside. Eggs undergo embryogenic development and second stage juveniles remain coiled inside the egg shell. Cysts detach easily from the roots and persist in the soil. Eggs can remain viable in cysts for several years (5-10).

The formation of syncytial cells is a prerequisite for nematode development these cells number only a few (2-3) and are small 5-6 days after nematode invasion, but increase in number (7-10) and grow 20 days later, thus occupying nearly the entire vascular area. This results in a disruption of xylem and phloem elements and an impairment of root function. The distal portion of the root may also die.

Sexual dimorphism is evident in the fourth stage and is pronounced in adults. Juvenile females remain sedentary as they moult, becoming more and more swollen until they are lemon-shaped. Fourth stage, or adult females rupture the root cortex and become externally visible. Males are sedentary and endoparasitic until the fourth juvenile stage and then become free living in the soil as adults.

Eggs do not become dormant or go through a stage of diapause (Greco et al., 1992b). The highest rate of hatching occurs when the temperature is 15-25°C, whilst hatching is nil at 5 or 30°C and negligible at 10°C. A few juveniles can emerge from eggs contained inside females. Penetration of second stage juveniles within chickpea roots occurs equally well at 10 and 25°C. At 8°C juveniles also invade the roots but they do not develop further. Minimum basal temperature for development is 10°C, while at 30°C root invasion by juveniles is suppressed ( Kaloshian et al., 1986a ).

At 20°C, the day degrees required to reach different developmental stages within roots of chickpea are 80, 120, 180, 190 and 370 for third and fourth stage juveniles, adults, eggs and cyst formation, respectively. First and second stage juveniles are visible within the eggs after 15 and 17 days from their formation, respectively ( Kaloshian et al., 1986b ).

Under field conditions in Syria, females can be seen by mid March or April on the roots of winter and spring-sown chickpea, respectively. Cysts develop by early or late April. By harvest (early-mid June), most of the eggs within cysts are embrionated (Greco et al., 1988). On the Anatolian Plateau females and cysts can be found in July. Maximum nematode reproduction rates of about 250 and 75 on winter and spring chickpea, respectively, have been observed in microplots (Greco et al., 1988 1993) and of 17, after an initial population density of 2 eggs/g soil under field conditions (Saxena et al., 1992). A decline in the number of nematode eggs in the soil in the range of 33-50% was observed after 7 months in the absence of host plants. Generally, the nematode develops only one generation on spring chickpea, while there is evidence of a second generation, comprising a small proportion of the offspring of the first generation which occurs on winter chickpea and lentil sown in fields infested with


In conclusion, we have undertaken the first transcriptome analysis of the early parasitic J2 stages of H. avenae during incompatible infection, and identified some genes that may be important in plant parasitism. Additionally, a comparative analysis of gene expression between the pre-parasitic J2s and the early parasitic J2s provides additional information on some pivotal genes that are likely to be involved either in parasitism or nematode metabolism. Although the available unigene set is not complete enough to analyze the whole life cycle of incompatible infection, it still serves as sequences and functions integrated resource for further analysis and identification of some pivotal parasitism genes and specific regulatory pathways during early CCN invasion, and there will benefit better in-depth understanding the molecular mechanisms of CCN parasitism. Moreover, it also provided candidates for improving the CCN resistance in crops by engineering techniques. Functional validation of individual candidates and their respective roles in CCN parasitism are needed to be further investigated.

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