Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Cirsium arvense
(creeping thistle)

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Datasheet

Cirsium arvense (creeping thistle)

Summary

  • Last modified
  • 27 September 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Natural Enemy
  • Host Plant
  • Preferred Scientific Name
  • Cirsium arvense
  • Preferred Common Name
  • creeping thistle
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Plantae
  •     Phylum: Spermatophyta
  •       Subphylum: Angiospermae
  •         Class: Dicotyledonae
  • Summary of Invasiveness
  • Invasive characteristics include the ability of C. arvense to produce large numbers seeds, (up to 5,300 per plant: Hay, 1937),...

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Pictures

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PictureTitleCaptionCopyright
Leaves generally oblong in outline, margin variable from entire to deeply pinnately segmented, spiny. Male heads globular, somewhat smaller than the flask-shaped female heads.
TitleInflorescence and leaves - colour illustration
CaptionLeaves generally oblong in outline, margin variable from entire to deeply pinnately segmented, spiny. Male heads globular, somewhat smaller than the flask-shaped female heads.
CopyrightNOVARTIS
Leaves generally oblong in outline, margin variable from entire to deeply pinnately segmented, spiny. Male heads globular, somewhat smaller than the flask-shaped female heads.
Inflorescence and leaves - colour illustrationLeaves generally oblong in outline, margin variable from entire to deeply pinnately segmented, spiny. Male heads globular, somewhat smaller than the flask-shaped female heads.NOVARTIS
Stems 30-150 cm tall, slender, green, freely branched; leaves alternate, base sessile and clasping or shortly decurrent.
TitleShoots, leaves and stems
CaptionStems 30-150 cm tall, slender, green, freely branched; leaves alternate, base sessile and clasping or shortly decurrent.
CopyrightUSDA
Stems 30-150 cm tall, slender, green, freely branched; leaves alternate, base sessile and clasping or shortly decurrent.
Shoots, leaves and stemsStems 30-150 cm tall, slender, green, freely branched; leaves alternate, base sessile and clasping or shortly decurrent.USDA
C. arvense rosettes in a well-established patch in the autumn.
TitleRosettes
CaptionC. arvense rosettes in a well-established patch in the autumn.
CopyrightUSDA
C. arvense rosettes in a well-established patch in the autumn.
RosettesC. arvense rosettes in a well-established patch in the autumn.USDA
Florets all tubular, rose-purple to pinkish, less commonly white. Florets of female heads 23-26 mm long; tube 20-23 mm.
TitleFlowers
CaptionFlorets all tubular, rose-purple to pinkish, less commonly white. Florets of female heads 23-26 mm long; tube 20-23 mm.
CopyrightUSDA
Florets all tubular, rose-purple to pinkish, less commonly white. Florets of female heads 23-26 mm long; tube 20-23 mm.
FlowersFlorets all tubular, rose-purple to pinkish, less commonly white. Florets of female heads 23-26 mm long; tube 20-23 mm.USDA
Perennial herb spreading rapidly by horizontal roots (left) which give rise to aerial shoots (right).
TitleRoots and adventitious shoots
CaptionPerennial herb spreading rapidly by horizontal roots (left) which give rise to aerial shoots (right).
CopyrightUSDA
Perennial herb spreading rapidly by horizontal roots (left) which give rise to aerial shoots (right).
Roots and adventitious shootsPerennial herb spreading rapidly by horizontal roots (left) which give rise to aerial shoots (right). USDA
C. arvense patch in maize, with pappus on seedheads visible.
TitleCompetition in maize
CaptionC. arvense patch in maize, with pappus on seedheads visible.
CopyrightUSDA
C. arvense patch in maize, with pappus on seedheads visible.
Competition in maizeC. arvense patch in maize, with pappus on seedheads visible.USDA
C. arvense shoots with pappus on seedheads in windrowed spring wheat before combining.
TitleAffected wheat harvest
CaptionC. arvense shoots with pappus on seedheads in windrowed spring wheat before combining.
CopyrightUSDA
C. arvense shoots with pappus on seedheads in windrowed spring wheat before combining.
Affected wheat harvestC. arvense shoots with pappus on seedheads in windrowed spring wheat before combining.USDA
Aceria anthocoptes (rust mite); here on creeping thistle (Cirsium arvense), may have potential as a biological control agent of this weed. Note scale.
TitleNatural enemy
CaptionAceria anthocoptes (rust mite); here on creeping thistle (Cirsium arvense), may have potential as a biological control agent of this weed. Note scale.
CopyrightPublic Domain - Released by the United States Dept. of Agriculture/USDA-ARS/original image by Eric Erbe and digital colorization by Chris Pooley EMU
Aceria anthocoptes (rust mite); here on creeping thistle (Cirsium arvense), may have potential as a biological control agent of this weed. Note scale.
Natural enemyAceria anthocoptes (rust mite); here on creeping thistle (Cirsium arvense), may have potential as a biological control agent of this weed. Note scale.Public Domain - Released by the United States Dept. of Agriculture/USDA-ARS/original image by Eric Erbe and digital colorization by Chris Pooley EMU

Identity

Top of page

Preferred Scientific Name

  • Cirsium arvense (L.) Scop. (1772)

Preferred Common Name

  • creeping thistle

Other Scientific Names

  • Cirsium incanum Bieb.
  • Cirsium lanatum Spreng.
  • Cirsium setosum (Willd.) Bieb.
  • Cnicus arvensis Hoffm.

International Common Names

  • English: California thistle; Canada thistle; field thistle
  • Spanish: cardo
  • French: chardon des champs; cirse des champs; sarrette des champs
  • Portuguese: cardo-das-vinhas

Local Common Names

  • Denmark: ager-tidsel; mark-tidsel
  • Finland: pelto-ohdake
  • Germany: Ackerdistel; Acker-Kratzdistel; Feldkratzdistel
  • Italy: scardaccione; stoppione
  • Japan: ezonokitsuneazami
  • Netherlands: akkervederdistel
  • South Africa: Kanadese dissel
  • Sweden: akertistel
  • Yugoslavia (Serbia and Montenegro): palamida

EPPO code

  • CIRAR (Cirsium arvense)

Summary of Invasiveness

Top of page Invasive characteristics include the ability of C. arvense to produce large numbers seeds, (up to 5,300 per plant: Hay, 1937), spread through clonal propagation, and to produce allelopathic effects, all of which promote a wide distribution in agricultural landscapes (Kazinczi et al., 2001; Eber and Brandl, 2003). Crawley et al. (1999) listed C. arvense as among the seven most invasive of weeds in grasslands in the UK. Reproduction by seed chiefly contributes to dispersal, not persistent seed banks (Hill et al. 1989; Heimann and Cussans, 1996; Bond and Turner, 2003). C. arvense thrives in disturbed habitats, and is spread by ploughing and superficial cultivation. Root fragments spread by these means may produce new plants with fragments as small as 3 mm (Drlik et al., 2000; Stolcova, 2002). Larger fragments (21 cm) produced more vigorous shoots than smaller ones (5 cm) (Gustavsson, 1997). Roots readily withstand freezing, thawing and drying, and seeds may remain viable for up to 20 years of storage in the soil (Holm et al., 1991). In North America, C. arvense is on 33 of 38 possible noxious weed lists (Skinner et al., 2000).

Taxonomic Tree

Top of page
  • Domain: Eukaryota
  •     Kingdom: Plantae
  •         Phylum: Spermatophyta
  •             Subphylum: Angiospermae
  •                 Class: Dicotyledonae
  •                     Order: Asterales
  •                         Family: Asteraceae
  •                             Genus: Cirsium
  •                                 Species: Cirsium arvense

Notes on Taxonomy and Nomenclature

Top of page The taxonomy and plant description of C. arvense (L.) Scop. have been reported in several sources (Moore and Frankton, 1974; Moore, 1975; Holm et al., 1977; Donald, 1990), which include keys for the subspecies (varieties) (Moore and Frankton, 1974; Moore, 1975; Donald, 1990). C. arvense has the following varieties or subspecies: vestitum; integrifolium; arvense (syn. mite); horridum (all Wimm. & Grab. 1829). Variability in growth characteristics of different ecotypes has also been reviewed (Donald, 1990).

Description

Top of page (After Moore, 1975: pp 1033-1034.)

Perennial herb spreading rapidly by horizontal roots which give rise to aerial shoots. Stems 30-150 cm tall, slender, green, freely branched. Leaves alternate, the base sessile and clasping or shortly decurrent; leaves generally oblong in outline, margin variable from entire to deeply pinnately segmented, spiny. Variation in leaf characters (texture, vestiture, segmentation, spinyness) is the basis for the varieties.

Plants dioecious, all heads of a plant either male or female. Flower heads numerous, 1-5 per branch, 15-25 mm high and 1/3 to 1/4 as wide; male heads globular, somewhat smaller than the flask-shaped female heads. Involucre 10-20 mm high, outer phyllaries ovate, tough-textured, subulate-tipped (0.5 to 0.75-mm stout spine), surface and margins glabrous or lightly arachnoid and with a narrow glandular mid-line; inner phyllaries progressively longer, the innermost unarmed, apex flat, chartaceous, often purplish and erose. Florets all tubular, rose-purple to pinkish, less commonly white. Florets of female heads 23-26 mm long; tube 20-23 mm, lobes about 2 mm; the pistil well-developed but anthers vestigial or absent; florets of male heads 12-14 mm long, tube 7-8.5 mm, lobes 3-4 mm, pistil absent or present and superficially normal but with vestigial ovary; anther 4 mm long, pollen 42-44 µm diameter, tricolporate, exine spiny.

Pappus copious, white, feathery, 20-30 mm long on mature achenes; achenes 2.5-4 x 1 mm, straight or slightly curved, straw or light-brown.

The chromosome number 2n=34 has been reported for all the varieties, in both Europe and Canada.

Plant Type

Top of page Broadleaved
Herbaceous
Perennial
Seed propagated
Vegetatively propagated

Distribution

Top of page The introduction, spread, and distribution of C. arvense have been reviewed, including information gleaned from herbarium samples, perceptual surveys by experts, and scientific surveys in fields (Moore, 1975; Holm et al., 1977; Donald, 1990; Van Acker et al., 2000).

Although the centre of origin of C. arvense is unknown, it probably was originally native to south-eastern Europe and the eastern Mediterranean area (Moore, 1975). However, it has been resident throughout Europe, western Asia, and northern Africa for some time (Moore, 1975; Holm et al., 1977; Holm et al., 1979). More recently C. arvense has spread throughout the temperate zones of South America, Africa, New Zealand, and Australia in the Southern Hemisphere (Holm et al., 1977), and has been found in North America since the 17th century (Moore, 1975). Its historical spread into New Zealand and Australia has been reviewed (Meadly, 1957), as has its distribution in Australia in the 1970s (Amor and Harris, 1974).

The range of C. arvense covers an estimated 9 770 000 km² in North America, extending 2090 km north to south and 4700 km east to west (Moore, 1975). The northern extent of C. arvense is 59°N in Canada and its southern limit is 40°N in the United States (Erickson, 1983). The most severe infestations in the United States occur in the northern half of the country (Hodgson, 1971).

Distribution Table

Top 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/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Asia

AfghanistanRestricted distributionNativeHolm et al., 1979; USDA-ARS, 2004; EPPO, 2014
ArmeniaPresentNativeHolm et al., 1979; USDA-ARS, 2004
AzerbaijanPresentNativeUSDA-ARS, 2004
ChinaRestricted distributionIntroducedHolm et al., 1979; Wan et al., 1996; EPPO, 2014
Georgia (Republic of)PresentNativeUSDA-ARS, 2004
IndiaPresentIntroducedHolm et al., 1979; Balyan et al., 2000
IranPresentNativeHolm et al., 1979; USDA-ARS, 2004
JapanRestricted distributionIntroducedHolm et al., 1979; EPPO, 2014
Korea, DPRPresentIntroducedHolm et al., 1979
Korea, Republic ofPresentIntroducedHolm et al., 1979; Kang et al., 1996
LebanonRestricted distributionHolm et al., 1979; EPPO, 2014
PakistanPresentIntroducedHolm et al., 1979
TurkeyPresentNative Invasive Holm et al., 1979; Kaya and Zengin, 2000; USDA-ARS, 2004
TurkmenistanPresentNativeUSDA-ARS, 2004

Africa

AngolaPresentIntroducedHolm et al., 1979
South AfricaPresentIntroducedMoore, 1975; Holm et al., 1979
SudanPresentNativeHolm et al., 1979
SwazilandPresentIntroducedHolm et al., 1979
TunisiaPresentNativeHolm et al., 1979
ZimbabweRestricted distributionEPPO, 2014

North America

CanadaRestricted distributionIntroduced Invasive Darbyshire, 2003; EPPO, 2014
-AlbertaPresentIntroduced Invasive Moore, 1975; Darbyshire, 2003
-British ColumbiaPresentIntroduced Invasive Moore, 1975; Queen's Printer, 2001; Darbyshire, 2003
-ManitobaPresentIntroduced Invasive Moore, 1975; Darbyshire, 2003
-New BrunswickPresentIntroducedMoore, 1975; Darbyshire, 2003
-Newfoundland and LabradorPresentIntroducedMoore, 1975; Darbyshire, 2003
-Northwest TerritoriesPresentIntroducedDarbyshire, 2003
-Nova ScotiaPresentIntroduced Invasive Moore, 1975; Darbyshire, 2003
-OntarioPresentIntroduced Invasive Moore, 1975; Cowbrough, 2003; Darbyshire, 2003
-Prince Edward IslandPresentIntroducedMoore, 1975; Darbyshire, 2003
-QuebecPresentIntroduced Invasive Moore, 1975; Darbyshire, 2003
-SaskatchewanPresentIntroduced Invasive Moore, 1975; Queen's Printer, 1999; Darbyshire, 2003
-Yukon TerritoryPresentIntroducedDarbyshire, 2003
MexicoPresentIntroducedHolm et al., 1979
USAPresentIntroduced Invasive Hodgson, 1971
-AlaskaPresentIntroducedUSDA-NRCS, 2004
-ArizonaPresentIntroduced Invasive USDA-NRCS, 2004
-ArkansasPresentIntroducedUSDA-NRCS, 2004
-CaliforniaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-ColoradoPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-ConnecticutPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-DelawarePresentIntroduced Invasive USDA-NRCS, 2004
-HawaiiPresentIntroduced Invasive USDA-NRCS, 2004
-IdahoPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-IllinoisPresentIntroduced Invasive Hodgson, 1971
-IndianaPresentIntroduced Invasive Hodgson, 1971
-IowaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-KansasPresentIntroduced Invasive USDA-NRCS, 2004
-KentuckyPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-MainePresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-MarylandPresentIntroduced Invasive USDA-NRCS, 2004
-MassachusettsPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-MichiganPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-MinnesotaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-MissouriPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-MontanaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-NebraskaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-NevadaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-New HampshirePresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-New JerseyPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-New MexicoPresentIntroduced Invasive USDA-NRCS, 2004
-New YorkPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-North CarolinaPresentIntroduced Invasive USDA-NRCS, 2004
-North DakotaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-OhioPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-OklahomaPresentIntroduced Invasive USDA-NRCS, 2004
-OregonPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-PennsylvaniaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-Rhode IslandPresentIntroducedUSDA-NRCS, 2004
-South DakotaPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-TennesseePresentIntroducedUSDA-NRCS, 2004
-UtahPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-VermontPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-VirginiaPresentIntroducedUSDA-NRCS, 2004
-WashingtonPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-West VirginiaPresentIntroducedHodgson, 1971; USDA-NRCS, 2004
-WisconsinPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004
-WyomingPresentIntroduced Invasive Hodgson, 1971; USDA-NRCS, 2004

South America

ChileRestricted distributionIntroducedHolm et al., 1979; EPPO, 2014

Europe

AlbaniaPresentNativeUSDA-ARS, 2004
AustriaPresentNativeUSDA-ARS, 2004
BelarusPresentNative Invasive USDA-ARS, 2004
BelgiumRestricted distributionNativeHolm et al., 1979; USDA-ARS, 2004; EPPO, 2014
BulgariaWidespreadNative****Holm et al., 1979; EPPO, 2014
CroatiaPresentNativeStefanic et al., 1999
Czech RepublicWidespreadNativeHolm et al., 1979; Burysková, 1997; EPPO, 2014
Czechoslovakia (former)WidespreadNative****Holm et al., 1979; USDA-ARS, 2004; EPPO, 2014
DenmarkPresentNativeHolm et al., 1979; USDA-ARS, 2004
EstoniaPresentNative Invasive Kuill et al., 1999
FinlandRestricted distributionNativeHolm et al., 1979; Jutila, 1996; EPPO, 2014
FranceRestricted distributionNativeHolm et al., 1979; Barralis et al., 1996; USDA-ARS, 2004; EPPO, 2014
GermanyPresentNative Invasive Holm et al., 1979; USDA-ARS, 2004
GreecePresentNativeHolm et al., 1979
HungaryPresentNativeRadványi and József, 2000; USDA-ARS, 2004
IcelandPresentNativeHolm et al., 1979
IrelandPresentNativeUSDA-ARS, 2004
ItalyPresentNativeHolm et al., 1979; Rapparini, 1999; USDA-ARS, 2004
LatviaPresentNative Invasive Lapinsh et al., 2000; USDA-ARS, 2004
LithuaniaPresentNativeCiuberkis and Petraitis, 1998; USDA-ARS, 2004
MoldovaPresentNativeUSDA-ARS, 2004
NetherlandsPresentHolm et al., 1979
NorwayPresentNativeHolm et al., 1979; USDA-ARS, 2004
PolandPresentNativeHolm et al., 1979; Skrzyczynska, 1999
PortugalRestricted distributionNativeHolm et al., 1979; USDA-ARS, 2004; EPPO, 2014
RomaniaPresentNativeHolm et al., 1979; USDA-ARS, 2004
Russian FederationPresentNativeHolm et al., 1979; Bystraya et al., 2000; USDA-ARS, 2004
SlovakiaPresentNativeHolm et al., 1979; Danadová, 2000
SloveniaPresentNativeHolm et al., 1979
SpainPresentNativeHolm et al., 1979; Lete et al., 1997; USDA-ARS, 2004
SwedenPresentNative Invasive Holm et al., 1979; Gustavsson, 1994; USDA-ARS, 2004
SwitzerlandPresentNative Invasive Bacher et al., 1997; USDA-ARS, 2004
UKPresentNative Invasive Holm et al., 1979; Edwards et al., 2000; USDA-ARS, 2004
UkrainePresentNativeHolm et al., 1979; USDA-ARS, 2004
Yugoslavia (former)PresentNativeHolm et al., 1979; USDA-ARS, 2004
Yugoslavia (Serbia and Montenegro)PresentNativeHolm et al., 1979; Stefanovic et al., 1998; USDA-ARS, 2004

Oceania

AustraliaRestricted distributionIntroducedHolm et al., 1979; Australian Weeds Committee, 2004; EPPO, 2014
New ZealandPresentIntroduced Invasive Holm et al., 1979; Jessep, 1997

History of Introduction and Spread

Top of page C. arvense was recognized as a troublesome weed in its native region of southeastern Europe by the 16th century, and by the mid-18th century was common throughout Europe (Moore, 1975). C. arvense is thought to have been introduced to North America sometime in the 17th century (Nuzzo, 2000; Moore, 1975). It was brought over from Europe accidentally as a contaminant of farm seed in both New France and New England (Moore, 1975; Whitson et al., 2001). It quickly spread in these spread regions becoming a troublesome agricultural weed. Legislation was enacted as early as 1795 in Vermont and 1831 in New York to control C. arvense (Moore, 1975; Nuzzo, 2000). Presently, C. arvense is widespread throughout the temperate regions of North America between 37° to 59° N (Nuzzo, 2000; Moore, 1975).

Risk of Introduction

Top of page C. arvense has the potential to spread to new areas via contaminated crop seed, irrigation channels, or living organisms. There is very little chance that C. arvense will be spread deliberately as it is widely regarded as a pest in most temperate regions and has no value as an ornamental plant.

Habitat

Top of page C. arvense can infest many temperate agricultural crops (Moore, 1975; Donald, 1990; Donald and Nalewaja, 1990; Fay, 1990; Hunter et al., 1990). It is found in both disturbed (tilled) and no-tillage agricultural fields used for producing most annual, winter annual, and perennial agronomic and horticultural crops, as well as adjacent sites, including non-cropped undisturbed roadsides.

Habitat List

Top of page
CategorySub-CategoryHabitatPresenceStatus
Terrestrial
Terrestrial – ManagedCultivated / agricultural land Present, no further details Harmful (pest or invasive)
Managed forests, plantations and orchards Present, no further details Harmful (pest or invasive)
Managed grasslands (grazing systems) Present, no further details Harmful (pest or invasive)
Disturbed areas Present, no further details Harmful (pest or invasive)
Rail / roadsides Present, no further details Harmful (pest or invasive)
Urban / peri-urban areas Present, no further details Harmful (pest or invasive)
Terrestrial ‑ Natural / Semi-naturalNatural forests Present, no further details
Natural grasslands Present, no further details Harmful (pest or invasive)
Riverbanks Present, no further details

Hosts/Species Affected

Top of page C. arvense is perennial and often considered 'noxious' (apearing on 33 noxious lists in North America) (Skinner et al., 2000), it has thus been of concern to farmers who grow cereals, oilseeds and forage products. It has been recorded infesting more than 27 crops in 37 countries (Moore, 1975; Holm et al., 1977).

C. arvense can infest many temperate agricultural crops (Moore, 1975; Donald, 1990; Donald and Nalewaja, 1990; Fay, 1990; Hunter et al., 1990). It is found in both disturbed (tilled) and no-tillage agricultural fields used for producing most annual, winter annual, and perennial agronomic and horticultural crops, as well as adjacent sites, including non-cropped undisturbed roadsides, riverbanks, forest edges and open meadows. C. arvense may also be a problem for the propagation of trees (Clay and Dixon, 1997; Siipilehto, 2001).

Host Plants and Other Plants Affected

Top of page
Plant nameFamilyContext
Aeschynomene falcata (joint vetch)FabaceaeMain
Allium cepa (onion)LiliaceaeOther
Allium porrum (leek)LiliaceaeOther
Allium sativum (garlic)LiliaceaeOther
Apium graveolens (celery)ApiaceaeOther
Arachis hypogaea (groundnut)FabaceaeOther
Asparagus officinalis (asparagus)LiliaceaeOther
Avena sativa (oats)PoaceaeMain
Beta vulgaris var. saccharifera (sugarbeet)ChenopodiaceaeOther
Brassica juncea var. juncea (Indian mustard)BrassicaceaeOther
Brassica napus var. napus (rape)BrassicaceaeMain
Brassica nigra (black mustard)BrassicaceaeOther
Brassica oleracea (cabbages, cauliflowers)BrassicaceaeOther
Brassica oleracea var. botrytis (cauliflower)BrassicaceaeOther
Brassica oleracea var. italica (broccoli)BrassicaceaeOther
Brassica oleracea var. viridis (collards)BrassicaceaeOther
Brassica rapa subsp. rapa (turnip)BrassicaceaeOther
Bromus (bromegrasses)PoaceaeMain
Capsicum annuum (bell pepper)SolanaceaeOther
Carthamus tinctorius (safflower)AsteraceaeMain
Chamomilla recutita (common chamomile)AsteraceaeOther
Cicer arietinum (chickpea)FabaceaeOther
Citrullus lanatus (watermelon)CucurbitaceaeOther
Cucumis melo (melon)CucurbitaceaeOther
Cucumis sativus (cucumber)CucurbitaceaeOther
Cucurbita maxima (giant pumpkin)CucurbitaceaeOther
Cucurbita pepo (marrow)CucurbitaceaeOther
Dactylis glomerata (cocksfoot)PoaceaeMain
Daucus carota (carrot)ApiaceaeOther
Fagopyrum esculentum (buckwheat)Main
Fragaria ananassa (strawberry)RosaceaeOther
Glycine max (soyabean)FabaceaeMain
Gossypium (cotton)MalvaceaeMain
Helianthus annuus (sunflower)AsteraceaeMain
Hordeum vulgare (barley)PoaceaeMain
Lactuca sativa (lettuce)AsteraceaeOther
Lagenaria siceraria (bottle gourd)CucurbitaceaeOther
Linum usitatissimum (flax)Main
Lotus corniculatus (bird's-foot trefoil)FabaceaeMain
Lupinus (lupins)FabaceaeMain
Malus domestica (apple)RosaceaeOther
Medicago sativa (lucerne)FabaceaeMain
Nicotiana tabacum (tobacco)SolanaceaeOther
Onobrychis viciifolia (sainfoin)FabaceaeMain
Panicum miliaceum (millet)PoaceaeMain
Petroselinum crispum (parsley)ApiaceaeOther
Phaseolus (beans)FabaceaeOther
Phaseolus lunatus (lima bean)FabaceaeOther
Phleum (timothies)PoaceaeMain
Piper nigrum (black pepper)PiperaceaeOther
Pisum sativum (pea)FabaceaeMain
Poa (meadow grass)PoaceaeMain
Polyphagous (polyphagous)Main
Prunus avium (sweet cherry)RosaceaeOther
Prunus domestica (plum)RosaceaeOther
Pyrus communis (European pear)RosaceaeOther
Raphanus sativus (radish)BrassicaceaeOther
Rubus fruticosus (blackberry)RosaceaeOther
Rubus idaeus (raspberry)RosaceaeOther
Secale cereale (rye)PoaceaeMain
Securigera varia (crown vetch)FabaceaeMain
Solanum lycopersicum (tomato)SolanaceaeOther
Solanum melongena (aubergine)SolanaceaeOther
Solanum tuberosum (potato)SolanaceaeOther
Sorghum bicolor (sorghum)PoaceaeMain
Spinacia oleracea (spinach)ChenopodiaceaeOther
Trifolium (clovers)FabaceaeMain
Triticum (wheat)PoaceaeMain
Triticum aestivum (wheat)PoaceaeMain
Vicia faba (faba bean)FabaceaeOther
Vigna angularis (adzuki bean)FabaceaeOther
Vitis vinifera (grapevine)VitaceaeOther
Zea mays (maize)PoaceaeMain

Biology and Ecology

Top of page C. arvense is a perennial broadleaved weed with an extensive, spreading perennial root system (Rogers, 1928; Hayden, 1934; Hodgson, 1971; Amor and Harris, 1974; Amor and Harris, 1975; Kigel and Koller, 1985). Adventitious root buds arise from its perennial roots to form new adventitious shoots (Hayden, 1934; Hamdoun, 1970a, 1970b, 1972). This is the major method of vegetative propagation of C. arvense after seedling establishment. Sexual reproduction is by seed (achenes).

C. arvense is adapted to temperate regions, those with moderate summer temperatures and moderate rainfall (450-900 mm/year) (Hodgson, 1968; Holm et al., 1977). The chief factors that limit its spread across continents have not yet been determined unambiguously. However, high summer temperatures may limit its southern spread in North America (Hoefer, 1981).

C. arvense grows on a broad range of soils, including acidic soils with pH levels between 3 and 4 (Dunsford et al., 1998) and soils with a variety of textures. C. arvense produced deeper perennial roots in clay or muck soils (3.8 mm deep) compared to sand, gravel (1 m deep), or limestone (1.8 m deep) soils (Detmers, 1927). Also, C. arvense must have some, as yet unquantified, tolerance to soil salinity, because it was found in 40% of non-marsh, dryland saline sites surveyed in Alberta, Canada (Braidek et al., 1984).

Sexual Reproduction

C. arvense seedlings have a juvenile vegetative period before established plants can flower in response to photoperiod (Bakker, 1960), but C. arvense can flower in the same growing season that seedlings emerge. The sexual reproductive system of C. arvense was reviewed by Kay (1985).

Moore (1975) stated that C. arvense was classified by plant taxonomists as dioecious [with male and female flowers on separate plants]. However, Hodgson (1968) observed that male plants of C. arvense occasionally produce achenes (termed 'seed' here), making the species 'imperfectly dioecious'. Lloyd and Mayall (1976) and Delannay (1977, 1979) verified Hodgson's (1968) observations. Seed produced by male plants were smaller and percentage germination was less than for seed formed by female plants.

The viability of C. arvense seed harvested at various times during seed formation and maturation was measured by Gill (1938). Seed germinated 0, 0, and 38% when seedheads were cut when flowering, in flower bud, and when fully mature, respectively (Kinch and Termunde, 1957; Derscheid and Schultz, 1960).

Seed Dispersal

C. arvense seed can be dispersed by transport in contaminated crop seed, feed, packing straw (Cox, 1913; Tonkin and Phillipson, 1973; Holm et al., 1977; Ball et al., 1982), and manure, as well as by irrigation water (Bruns and Rasmussen, 1953, 1957; Moore, 1975) and wind (McKay et al., 1959). Human-assisted dispersal has been more important for the worldwide spread of C. arvense than biological dispersal.

Seed Bank Levels and Seed Persistence

Early field research indicated that C. arvense seed could be quite persistent in soil. For example, some C. arvense seed germinated 22 years after burial 20 cm deep in the field (Madsen, 1962). Seed germinated 45-55% when unearthed after burial 25 cm deep for 5 years in the field (Kjaer, 1948) and 52% after burial 20 cm deep for 3 years (Dorph-Petersen, 1924). Thus, if C. arvense seed were buried > 20 cm deep by moldboard ploughing, a small proportion of seed would persist to re-infest soil if seed were subsequently unearthed by later tillage. However, these early seed persistence studies can be criticized because C. arvense seed were buried well below the depth of normal emergence.

Subsequent research on seed persistence, which simulated more realistic field conditions, showed that C. arvense seed were quite short-lived in the soil. These studies used shallow burial (2.5-7.5 cm deep) and periodic soil disturbance (Roberts and Chancellor, 1979), which are more typical of agricultural fields. Less than 1% of the total number sown remained after 2.5 years in Denmark (Bakker, 1960) and 5 years in England (Roberts and Chancellor, 1979). Most seed were lost from the soil seed bank by germination during the first year after burial at 7.5 cm (48-58%) or 2.5 cm (60%). Soil disturbance is known to speed the rate of loss of other weed seed from the soil seed bank (Roberts, 1964, 1981). When seed persistence of undisturbed seed was assessed in Canada (by emergence alone), no seedlings emerged 3 years after burial (Chepil, 1946). Likewise Hill et al. (1989) reported it was absent from the seed bank among different cropping systems. However, Eber and Brandl (2003) attributed regional persistence of C. arvense partially to the possession of persistent seed banks and Hakansson (2003) warned that C arvense occurring in agricultural headlands may re-infest fields.

Seedling Germination, Emergence, and Phenology in the Field

Almost all C. arvense seed can germinate at maturity and soon after dispersal (Moore, 1975). Fresh mature seed germinated well within 2 to 4 days after being excised from the pericarp (i.e. seed coverings) (Kay, 1985), suggesting that the pericarp restricted germination of freshly shed seed. Environmental effects on germination have been reviewed (Donald, 1994).

Some C. arvense seed can germinate on the soil surface, making the species 'pre-adapted' to reduced or zero tillage (Wilson, 1979). However, no C. arvense seedlings became established from seed spread on the soil surface of a mixed pasture in Australia (Amor and Harris, 1974). When seed were buried 0.5 to 1.0 cm deep, however, 6.8 to 12.6% of seed emerged (Amor and Harris, 1975).

With repeated monthly soil disturbance, C. arvense seedlings emerged from 68% of shallowly buried (2.5 cm deep) seed over 3 years (Roberts and Chancellor, 1979). As planting depth increased, seedling emergence decreased and seedlings from deeper-planted seed emerged later than more shallowly buried seed (Zilke and Derscheid, 1957). In contrast, Derscheid et al. (1956) reported that seed planted deeper than 1.3 cm failed to germinate, while soil type did not greatly influence emergence down to 2 cm (Bakker, 1960). When seed were buried at various depths as low as 6 cm in two sandy loam soils, emergence was greatest from 0.5 to 1.5 cm (Wilson, 1979).

In fields in England, few seedlings emerged in the first autumn after seed were sown in September (Roberts and Chancellor, 1979). Only 3 to 6 months were required for seed to become fully capable of germinating in the field. Winter environments usually prevent C. arvense seed germination. After seed maturation, primary dormancy was short-lived but was followed by longer-term, environmentally enforced (secondary) dormancy. Environmentally enforced dormancy develops if seed experience environmental conditions that prevent germination (Bakker, 1960).

Most seedlings emerged in April and May following burial in the previous autumn, but a small number persisted longer (Bakker, 1960). In Denmark, C. arvense seed were able to germinate 88% in the spring after autumn burial (Dorph-Petersen, 1924). In England, 60% of seed planted 2.5 cm deep and 48-58% of seed planted 7.3 cm deep emerged in the spring after autumn planting (Roberts and Chancellor, 1979).

C. arvense seedlings grow slowly at first and are poor competitors with low shade tolerance (Holm et al., 1977). Shading at 60-70% of full sunlight severely restricted shoot growth, and seedlings died at 80% shade (Bakker, 1960). Plants growing in forests were tall, spindly, and flowered less than plants growing in the open (Detmers, 1927). Clearly, C. arvense grows best under unshaded conditions.

Vegetative Growth of Established Shoots

Emergence phenology of adventitious shoots
Seasonal environment may restrict adventitious root bud elongation of C. arvense, rather than endogenous physiological dormancy (Grondal et al., 2003). In Nebraska (USA), when roots were periodically unearthed, brought indoors, and grown in a high-humidity incubator at 15°C for 2 weeks, there were no seasonal cycles of adventitious root bud activity (McAllister and Haderlie, 1985).

Hodgson (1968) observed that rosettes of C. arvense enlarged for 3 weeks before shoots began to grow vertically. Later-emerging shoots may elongate without rosette formation. Seasonal emergence of adventitious shoots in England (Lawson et al., 1974) and North America (Moore, 1975) has been summarized (Donald, 1994). A study of root longevity in sheep pastures in New Zealand concluded that very little of the overwintering biomass stored in C. arvense (6-10%) persisted into the next growing season (Bourdot et al., 2000).

C. arvense shoots can be propagated from lateral buds at internodes on stem segments (Magnusson et al., 1987) and from adventitious root buds arising on root segments (Moore, 1975). The relative importance of shoots arising from buried vertical stems versus adventitious root buds has not been determined in the field, although it is probably minor.

Adventitious shoot density
C. arvense often forms distinct circular or semicircular patches in fields. C. arvense shoots arising from adventitious root buds often exclude other species in the centre of patches (Bendall, 1975; Stachon and Zimdahl, 1980; Wilson, 1981b).

Typical C. arvense shoot densities in surveys of cereal fields have been summarized (Donald, 1994). Densities declined in the centres of 4- to 5-m-wide patches in Nebraska. In Australia, plant density and height were also reduced near the centre of a 28-m-wide patch from the patch borders, suggesting senescence (Amor and Harris, 1975). Earlier, Pavlychenko (1943) had observed that during a prolonged drought, dense patches of C. arvense became ring-like. Perhaps environmental stress rather than autotoxicity may limit the emergence of C. arvense in well-established patches, especially when patches are stressed by drought.

Under favourable conditions, a C. arvense patch can spread rapidly by vegetative means. One 7.5-cm-long cutting formed a solid patch 7.2 m wide after 3 years of uninterrupted growth (Pavlychenko et al., 1940). Patches spread laterally 0.8-1.6 m per year in Australian pastures, depending upon site and year (Amor and Harris, 1975), 6 m per year in the USA (Hayden, 1934), and 0.8-1.3 m per year in Europe (Bakker, 1960), depending upon site and year. Large patches tend to increase less rapidly than small ones, although established C. arvense clones tend to spread more rapidly (up to 12m per year: Chancellor, 1970) than recently formed clones (Amor and Harris, 1975; Eber and Brandl, 2003). Patches have been observed to degenerate behind an advancing front of C. arvense, likely due to autotoxicity (Amor and Harris, 1975; Bendall, 1975). The allelopathic effect of C. arvense on other plants also aids in formation of dense patches (Stachon and Zimdahl, 1980; Kazinczi et al., 2001) and was found to inhibit other weed species in a cropping situation (Hari et al., 2002).

Crop and land management can reduce the rate of C. arvense patch growth (Donald, 1990). For example, C. arvense did not spread as rapidly in grazed as in ungrazed pastures in Australia (Amor and Harris, 1975).

Roots

Development
After shallowly buried seed germinated, seedling roots penetrated 5 to 10 cm before shoots emerged (Bakker, 1960). Horizontal roots developed and adventitious shoots emerged within 6 to 8 weeks after establishment. In Canada, the first lateral roots were initiated at the two-leaf stage when seedings were 4 to 5 weeks old (Friesen, 1968). Adventitious root buds formed on the tap root (Sagar and Rawson, 1964; Friesen, 1968), but the depth of adventitious root bud formation varied. After new C. arvense seedlings became established, their ability to survive clipping by sending up adventitious shoots increased as plants aged (Wilson, 1979).

Depth
The depth distribution of C. arvense roots in the soil profile is controversial. In Montana, USA, most roots of 2-year-old plants were found 7.5-30 cm deep (Hodgson, 1968): 54, 30, and 16% of the roots extracted were found in 7.5-22.5, 22.5-37.5, and 37.5-52.5 cm depths, respectively.

Estimates of the maximum depths to which roots extend also vary. Arny (1932) observed that most roots grew no deeper than 40 cm, whereas most horizontal roots were found 20-30 cm deep. Vertical roots grew 1.8-3 m deep in Minnesota, USA (Arny, 1932) and 1.5-1.8 m deep in Iowa (Hayden, 1934). Soil type influenced rooting depth. Maximum depths of 1, 1.8, 3.8, and 4.5 m were observed in sand or gravel, limestone, muck soil, and clay soils, respectively (Detmers, 1927). Maximum depths of 2.4-2.7 m were reported elsewhere (Kiltz, 1930). A 10-year-old undisturbed stand in Canada had roots extending down to 2 m, but most roots were in the top 20 cm (Nadeau and VandenBorn, 1989). Restrictive soil conditions, such as hard pans, gravel or sand layers, alkaline or high calcium soil horizons, and high water tables, may restrict the maximum depth of root penetration (Rogers, 1928). Roots can grow along ped faces in the subsoil (W Donald, University of Missouri, USA, personal communication, 1995).

The maximum depth of emergence of adventitious shoots from adventitious root buds was reported as 0.9 m in Canada (Friesen, 1968).

When left undisturbed, a single C. arvense plant produced 26 emerged adventitious shoots, 154 underground adventitious root buds, and 111 m of roots (> 0.5 mm diameter) after 18 weeks of growth outdoors in boxes (Nadeau and VandenBorn, 1989). Year-to-year variation in root growth was observed (Nadeau and VandenBorn, 1990). Roots sampled down to 1.8 m produced an equivalent number of adventitious shoots compared to shallower sampling depths from a 10-year-old undisturbed stand.

Drought Tolerance

Although the perennial root system of C. arvense probably allows it to survive drought years better than annuals, biomass of perennial roots and numbers of adventitious root buds decrease after several years of drought (Donald, 1992). Drought can suppress subsequent shoot growth for 12 months (Amor and Harris, 1977). The density of C. arvense shoots and adventitious root buds growing in continuous spring wheat decreased in the year following drought years (Donald and Prato, 1992a, 1992b).

Response to Flooding

C. arvense can survive flooding, but this stress resulted in loss of leaves and reduced shoot and root growth (Rogers, 1928; Bakker, 1960; Lenssen et al., 1998).

Associations

C. arvense is associated with various animals that create soil disturbances (e.g. rabbits, moles), as well as grazing animals that reduce competing vegetation (Edwards et al., 2000). In northern Germany, an invasion of rats in permanent pastures aided the spread of C. arvense by housing the vegetative portions of the weed in their tunnels (Holm et al., 1991). Pemberton and Irving (1990) found that C. arvense seeds possess elaiosomes, suggesting a possible role for ants in seed dispersal. This mechanism (myrmechochory) may help C. arvense escape predation, avoid competition, or promote germination and establishment in favorable substrates (Pemberton and Irving, 1990). C. arvense populations are also associated with numerous insects, many of which feed on various parts of the plant (Moore, 1975; Eber and Brandl, 2003). Kruess (2003) observed increased insect abundance and diversity was associated with increased abundance of C. arvense on fields in Germany during fallow periods.

C. arvense has been found in almost every plant community in the temperature regions of the world. In a study in Colorado, the following species were associated with dense Canada thistle stands: American bullrush (Scirpus americanus), creeping bentgrass (Agrostis palustris), longstem spikerush (Eleocharis macrostachys), and saltgrass (Distichlis stricta) (Donald, 1994). In moderately dense Canada thistle stands the following species were associated with C. arvense: giant ragweed (Ambrosia trifida), horseweed (Conyza canadensis), common lambsquarters (Chenopodium album), redroot pigweed (Amaranthus retroflexus), prairie sunflower (Helianthus petiolaris), smooth dock (Rumex altissimus), and foxtail barley (Hordeum jubatum) (Donald, 1994). In agricultural fields, C. arvense tends to be associated with other similar and persistent perennial weeds.

Ecological succession

C. arvense is an early successional plant, and is typically found in disturbed areas as a part of the initial postdisturbance community. Once established, a single seedling can form a large patch of stems through vegetative propagation of the root system (Moore, 1975). C. arvense is commonly phased out of natural areas once competing plants create a canopy that inhibits the growth and spread of the weed (Hallgren, 1976; Edwards et al., 2000; Vulink et al., 2000; Stolcova, 2002). Rejmanek and Leps (1996) demonstrated that during a six-year period, the competitive relationship between C. arvense and the shrub Arctostaphylos patula shifted as the shrub gained in stature. A five-year study in an agricultural area in Bavaria recorded that 30% of patches became extinct each year, with most of these consisting of small patches subject to human management or intrinsic factors such as lack of suitability of the site for long-term establishment of C. arvense (Eber and Brandl, 2003). In contrast, mountain grazing lands in Poland experienced a marked increase in C. arvense populations four years after grazing had ceased (Kasperczyk and Szewczyk, 1999).

Plant genetics

C. arvense is capable of clonal reproduction from buds on horizontal roots (Lalonde and Roitberg, 1994; Whitson et al., 2001). Although clumps of C. arvense may appear uniform, DNA testing has demonstrated that several phenotypes may be present in a single clump (Heimann and Harding, 1996). Sexual reproduction by C. arvense was reviewed by Heimann and Cussans (1996). In terms of reproduction from seed, C. arvense is an obligate outcrosser, thus producing genetically diverse progeny (Lalonde and Roitberg, 1994; Pollak and Bailey, 2001). However, seed set may often be limited by availability of pollen in clonal populations of this normally dioecious plant, which frequently exhibits female-biased offspring sex ratios (Lalonde and Roitberg, 1994). Hermaphrodite flowers are occasionally produced (Heimann and Cussans, 1996). The chromosome number for all C. arvense varieties is 2n=34 (Nuzzo, 2000).

Air Temperature

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Parameter Lower limit Upper limit
Absolute minimum temperature (ºC) -35
Mean annual temperature (ºC) 1 20
Mean maximum temperature of hottest month (ºC) 22 32
Mean minimum temperature of coldest month (ºC) -14

Rainfall

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ParameterLower limitUpper limitDescription
Mean annual rainfall450900mm; lower/upper limits

Soil Tolerances

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Soil drainage

  • free
  • impeded

Soil reaction

  • acid
  • neutral

Soil texture

  • heavy
  • light
  • medium

Special soil tolerances

  • saline

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Aceria anthocoptes Herbivore
Aceria anthocoptes Herbivore
Alternaria cirsinoxia Pathogen
Altica carduorum Herbivore Leaves/Roots/Stems
Aphis fabae cirsiiacanthoidis Herbivore
Apion onopordi Herbivore
Brachycaudus cardui Herbivore
Capitophorus carduinus Herbivore
Cassida rubiginosa Herbivore Leaves/Roots/Stems Maryland; Virginia
Ceutorhynchus litura Herbivore Stems Montana; Saskatchewan; New Zealand
Cleonis pigra Herbivore Stems
Dasyneura gibsoni Herbivore
Erysiphe mayorii var. mayorii Pathogen
Larinus planus Herbivore
Lobesia abscisana Herbivore
Mycosphaerella cirsii-arvensis Pathogen Leaves/Stems
Orellia scorzonerae Herbivore
Phoma destructiva Pathogen
Phomopsis cirsii Pathogen
Pseudomonas syringae pv. tagetis Pathogen
Puccinia punctiformis Pathogen Leaves
Puccinia suaveolens Pathogen
Ramularia cirsii Pathogen Leaves
Rhinocyllus conicus Herbivore Inflorescence
Sclerotinia sclerotiorum Pathogen Roots
Septoria cirsii Pathogen
Spaeroderma testaceum Herbivore
Tephritis cometa Herbivore Inflorescence
Terellia ruficauda Herbivore Inflorescence
Terellia ruficauda Herbivore
Uroleucon cirsii Herbivore
Urophora aprica Herbivore
Urophora cardui Herbivore Stems New Zealand; Ontario; Saskatchewan
Urophora eriolepidis Herbivore
Urophora solstitialis Herbivore
Urophora stylata Herbivore
Verticillium dahliae Pathogen
Xyphosia miliaria Herbivore Inflorescence

Notes on Natural Enemies

Top of page Zwolfer (1965) studied the natural enemies of C. arvense in its native range in Europe. Moore (1975) listed arthropods that were observed to feed on C. arvense in North America, where it is an introduced weed. Relatively recently, Perju et al. (1995) collected 61 insect species on C. arvense in Romania, 25 of which were clearly attacking parts of the plant; further work is needed to better characterize this list, but the study indicates potential for the identification of new agents. Wherever C. arvense has been introduced, native organisms have colonized it but do not control it. A biological control programme has been under development in New Zealand for sometime (Cameron et al., 1989; Julien, 1992). By 1997, candidate species were narrowed to four: Lema cyanella, Altica carduorum, Ceutorhynchus litura and Urophora cardui (Jessep, 1997). Similar work has been undertaken in other parts of the world (e.g. North America and China).

There are limitations to the adoption of some of the natural enemies listed as biological control agents against C. arvense. For example, Altica carduorum, a chrysomelid beetle that defoliates and feeds on floral buds in Europe, failed to overwinter in northern USA and Canada (Peschken, 1981; Trumble and Kok, 1982). A population of A. carduorum was recently characterized from a similar climatic region in China, and a phenological model indicated that this strain of A. carduorum had the potential to overlap the range of C. arvense on the Canadian prairies (Lactin et al., 1997). However, its ability to develop on other North American species of Cirsium apparently precludes its use in North America (Wan and Harris, 1997). Cassida rubiginosa defoliated young plants by more than 50%, but did not damage old plants. It is naturalized in eastern Canada, but was slow to establish in Montana (Watson and Keogh, 1980; Peschken, 1981; Forsyth and Watson, 1985; Monnig, 1987), may suffer high winter mortality (Spring and Kok, 1999) and is most damaging at higher temperatures (Bacher and Schwab, 2000).

Ceuthorhynchus litura is a stem-boring weevil that mines leaf veins and root crowns. It may also transmit Puccinia punctiformis, a fungus that causes rust on C. arvense. C. litura was introduced into Canada and the USA from Europe for biological control; however, it has a limited dispersal ability (Peschken and Beecher, 1973; Trumble and Kok, 1982; Rees, 1990). Another European weevil, Cleonis pigra, also feeds at the root crown: feeding leads to gall formation. However, C. pigra does not control C. arvense because the death of infected shoots is limited (Watson and Keogh, 1980; Forsyth and Watson, 1985).

The aphid Brachycaudus cardui, which feeds on floral buds in the USA, is not thought to be a good biological control agent (Detmers, 1927).

Urophora cardui, a European tephritid which forms stem galls, was released in Canada and the USA but only had a slight impact on C. arvense (reduced shoot height) (Watson and Keogh, 1980; Peschken et al., 1982). Another tephritid, Terellia ruficauda, had little effect on established stands in pastures in Canada (Southey and Staniland, 1950; Watson and Keogh, 1980).
A mite, Aceria anthocoptes, specific to C. arvense, was discovered occurring on C. arvense throughout the mid-eastern USA (Ochoa et al., 2001). A. anthocoptes and A. leonthodontis occur in Europe (Petanovic et al., 1997).

Means of Movement and Dispersal

Top of page Natural dispersal

It is thought that the primary means of long distance travel for C. arvense seeds are dispersal by wind and water, transporting the plumed seeds to new locales (Holm et al., 1991). However, studies have shown that many of the plumes break off and drift away while the seeds remain in the head (Holm et al., 1991).

Viable seeds of C. arvense have been recovered from irrigation water after prolonged submersion (Donald, 1994; Bond and Turner, 2003). After 2 and 22 months of submersion in irrigation water, seeds germinated at a rate of 54 to 92% and 19%, respectively (Donald, 1994). Seeds have also been observed in surface water drainage areas (Drlik et al., 2000).

Vectors

Seeds can be carried to new locations by sticking to the clothes and shoes of humans, and to the fur and feet of animals (Holm, 1991). Birds that feed on the seeds of C. arvense may carry the plant to new locations. Holmes and Froud-Williams (2001) found that some of the C. arvense seeds ingested by chaffinches were still able to germinate. Rodents that collect and store the vegetative portions of C. arvense have resulted in the appearance of C. arvense in areas that previously did not have the weed (Bond and Turner, 2003). C. arvense may use ants as a means of seed dispersal. Seeds of C. arvense provide a fat- and protein-rich elaiosomes for the ant, and in return the ant carries the seeds away from the parent plants and into or near the ant nest (Pemberton and Irving, 1990). Invasion also occurs via roadways (e.g. Sudbrink et al., 2001).

Agricultural practices

The movement of C. arvense seeds via agricultural practices is primarily responsible for spreading C. arvense between different regions (Donald, 1994). Seeds of C. arvense have been a contaminant of various crops seeds, resulting in the unintentional spread of the weed to new locations that were previously not effected (Bond and Turner, 2003). As well, the seeds have been found in packing hay and feed for livestock (Holm et al., 1991; Donald, 1994). The animals can then carry the seeds externally or internally, as viable seeds have been found in the manure of heifer and dairy herds (Mt Pleasant and Schlather, 1994).

The spread of C. arvense via vegetative propagation after initial establishment is more important for local spread (Donald, 1994). Common agricultural practices such as ploughing and cultivation distribute fragments of stems and roots allowing the plant to become further spread (Holm et al., 1991; Stolcova, 2002). Root fragments have the ability to survive adversity and regenerate from small pieces. Root fragments as small as 3 - 6 mm are able to produce shoots, and shoot fragments as small as 6 cm were able to produce new shoots (Hayden, 1934). C. arvense also benefits from fields that are left fallow following cultivation or grazing (Kruess, 2001; Kasperczyk and Szewczyk, 1999). Although seeds from C. arvense plants in field margins may result in re-invasion of fields (Hakansson, 2003), the risk appears relatively low (Blumenthal and Jordan, 2001). Another potential source of invasion is the use of agricultural practices that allow survival of large populations of C. arvense, such as in the case in areas with "low levels of agricultural technology" (Reintam and Kuht, 1999).


Movement in Trade

C. arvense seed has been introduced to new locations as an impurity in seed stocks of small grains, as well as in packing straw (Holm et al., 1991).

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Land vehiclesFarm vehicles Yes
Plants or parts of plantsContaminated crop seed Yes
Soil, sand and gravel Yes

Plant Trade

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Plant parts not known to carry the pest in trade/transport
Bark
Bulbs/Tubers/Corms/Rhizomes
Flowers/Inflorescences/Cones/Calyx
Fruits (inc. pods)
Growing medium accompanying plants
Leaves
Roots
Seedlings/Micropropagated plants
Stems (above ground)/Shoots/Trunks/Branches
True seeds (inc. grain)
Wood

Impact Summary

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CategoryImpact
Animal/plant collections None
Animal/plant products Negative
Biodiversity (generally) Negative
Crop production Negative
Environment (generally) Negative
Fisheries / aquaculture None
Forestry production None
Human health None
Livestock production Negative
Native fauna None
Native flora Negative
Rare/protected species Negative
Trade/international relations None
Transport/travel None

Impact

Top of page C. arvense is a major pest and is considered one of the world's worst weeds, ranked as the third most important weed in Europe (Friedli and Bacher, 2001). Yield losses due to C. arvense occur in horticultural crops, field crops, pastures, rangelands, lawns, vineyards and orchards (Hodgson, 1964; 1968; Moore, 1975; Holm et al., 1977; Varadi et al., 1987). It is primarily a weed in perennial forage crops and pastures in North America and Europe and is considered a weed of 27 crops in 37 countries (Holm et al., 1991). C. arvense causes greater crop losses than any other broadleaf weed in its growth range, which includes 10 million km2 in Canada and the northern United States (Drlik et al., 2000). The agricultural crops it affects worldwide include barley, flax, millet, oats, rye, sorghum, wheat and other cereals, rape, canola, corn, beans, peas and other vegetables, vineyards, and orchards (Holm et al., 1991; Van Acker et al., 2000). In the Canadian prairies, a survey of 452 fields (212 wheat, 71 barley, 28 oat, 108 canola, 33 flax) found that C. arvense was the fourth most abundant weed with an average of 1.3 plants/m2. It was fifth most abundant in 1978-1981 and eighth most abundant in 1986 (Van Acker et al., 2000). It has been estimated that C. arvense causes annual losses in Canada of $3.6 million CAD in wheat alone (Peschken et al., 1980b).
C. arvense tends to form relatively isolated patches when it occurs in crops, except in reduced tillage or no-tillage farming systems. The increase in land under reduced tillage systems since the 1980s has lead to increased densities of C. arvense in a variety of cropping systems (Mayor and Maillard, 1995; Mills et al., 1997; Torresen et al., 2003). Although it does not 'take over' entire fields, it may require costly treatment to limit its spread. Donald and Khan (1996) found that among all yield components, the greatest reduction in spring wheat yield due to C. arvense was through early competition that reduced crop density. C. arvense is among the ten most frequently listed noxious weeds in North America (Skinner et al., 2000) and has thus been an important concern to farmers who grow cereals, oilseeds and forage products. Donald (1990) reviewed the economic impact of C. arvense on crop production and the yield loss.

The relative extent to which increasing densities of C. arvense reduce yield has been determined in tilled cropping systems for: winter and spring wheat (45 to 55% maximum yield loss) (Hodgson, 1955; Hodgson, 1968; Peschken et al., 1980b); barley (73% maximum yield loss) (Hodgson, 1955; O'Sullivan et al., 1982); oats (45% maximum yield loss) (Hodgson, 1955); Brassica napus (60% maximum yield loss) (O'Sullivan et al., 1985), alfalfa (Schreiber, 1967) and faba bean (8-12% maximum yield loss) (Kalburtji and Mamolos, 2001). Established natural infestations of C. arvense were used in the above studies (Hodgson, 1968; O'Sullivan et al., 1982, 1985), except for alfalfa (Schreiber, 1967) in which spaced transplants were used.
In pastures, C. arvense reduces forage consumption and cattle will not graze near the plants because of the sharp spines on its leaves (Holm et al., 1991; Drlik et al., 2000). Thus, it restricts the area available for livestock grazing (Donald, 1990; Edwards et al., 2000). A study found that 47% of New Zealand dairy farmers listed C. arvense as being a problem in their pastures (Bourdot et al., 1994). C. arvense can also have a detrimental effect on the health of livestock, as a study found that 16 of 150 weaned calves developed signs of polioencephalomalacia (Loneragan et al., 1998). There were signs of high H2S concentrations found in these calves rumen contents, which was a result of high C. arvense content in hay (Loneragan et al., 1998). C. arvense was recorded as an alternate host of Alfalfa mosaic virus in New Zealand (Fletcher, 2001) and Beet Necrotic Yellow Vein Virus (BNYVV) in Russia (Kutluk et al., 2000).
C. arvense has been investigated for use as an allelopathic agent, against either weeds or soil pathogens (Forleo, 2002; Hari et al., 2002). Extracts were also highly effective against the bacteria Xanthomonas campestris pv. oryzae [X. oryzae pv. oryzae] and Erwinia chrysanthemi pv. zeae (Kaushal-Gautam et al., 2001).

Environmental Impact

Top of page The natural communities where C. arvense has an impact include various non-forested plant communites such as prairies, barrens, savannas, glades, sand dunes, fields and open meadows that have been impacted by disturbance (White et al., 1993; Moore, 1975). It negatively impacts natural environments by crowding out and replacing native grasses and forbs, changing the structure and species composition of natural plant communities and reduces species diversity (White et al., 1993). Although primarily seen as a weed of field and horticultural crops or of natural areas, other environments are also affected such as turf, landscape and nurseries (Gao et al., 1999). As various countries put in place ecological compensation areas or set aside land formerly in agriculture, these lands become vulnerable to invading C. arvense which is often listed as the most common weed in these areas (Davies et al., 1994; Gustavsson, 1994; Barralis et al., 1995; Jewett et al., 1996; Bacher et al., 1997; Bacher and Schwab, 2000; Allen et al., 2001; Nemeth, 2001).

C. arvense produces allelochemicals that are released into the soil that can be toxic to surrounding vegetation, and the phytotoxicity of soil incorporated plant parts can persist for up to 9 weeks (Kazinczi et al., 2001). Kazinczy et al. (2001) found that root and foliage extracts reduced the radicle growth of barley, cucumber, green foxtail and redroot pigweed, and the germination of turnip, soybean, wheat, flax, barley, corn, alfalfa and sunflower. Annual weeds were reduced within stands of C. arvense, although some perennial grasses (Agrostis palustris, Distichlis stricta) and shrubs (Scirpus americanus and Eleocharis macrostachys) persisted (Stachon and Zimdahl, 1980). Species diversity decreased markedly from the periphery to the centre of C. arvense patches (Stachon and Zimdahl, 1980). Water extracts of roots or shoots or soil amended with dried C. arvense roots or shoots reduced the seed germination, establishment, and seedling growth of C. arvense (Donald, 1994).

C. arvense may act as a bioindicator of contamination and used to remediate soil subject to heavy metal pollution, and has been observed to accumulate lead, zinc and chromium (Samkaeva et al., 2001).

Impact: Biodiversity

Top of page The impact C. arvense has on natural areas is a relatively new area of study and very little is known on how the plant impacts such ecosystems and the biodiversity. Levine (2000) found in a California study that the most diverse plant communities were the most susceptible to invasive plants, including C. arvense. Negative impacts of C. arvense invasions on biodiversity include crowding out and replacing native grasses and forbs, decreasing the species diversity of an area, and altering ecosystem structure and composition (White et al., 1993). Enhancement of waterfowl populations may require control of C. arvense, to maintain plant community diversity (Krueger-Mangold et al., 2002). C. arvense is seen as a major threat to the Colorado butterfly plant (Gaura neomexicana subsp. coloradensis), listed by the US Fish and Wildlife Service as a threatened species (Munk et al., 2002). In Wyoming, C. arvense infestations have contributed to the elimination of the endangered Colorado butterfly plant (Cheater, 1992). In some contexts, C. arvense may also be beneficial to native organisms. In Washington, C. arvense provided cover for the endangered Columbian white-tailed deer in the summer, allowing deer to utilize previously unused areas (Suring and Vohs, 1979).

Nine hybrids between C. arvense and other Cirsium species were recorded in Europe (Hegi, 1929). Only one of the hybridizing species (C. palustre) has been introduced to North America (Moore, 1975). A possible hybrid between C. arvense and C. hookerianum (a Cirsium species native to North America) was described by Moore and Frankton (1965) as occurring in British Columbia, but did not appear to have significant impacts (Moore, 1975).

C. arvense has invaded several national parks and protected areas. It is one of the most prominent non-native plant species infesting four Great Lakes National Parks (Apostle Islands, Indiana Dunes, Pictured Rocks, and Sleeping Bear Dunes) (Bennett, 2001). In Yellowstone National Park in the USA, C. arvense invaded several burned areas from 1972 to 1988 and now occurs along horse trails, foot trails, and roadways (Turner et al., 1997). A study to determine the frequency of exotic plant species in Yellowstone National Park campgrounds found that C. arvense was the most frequent of the exotic plant species, being found in 6 of 11 campsites (Allen and Hansen, 1999). Following a fire in 1988, several burned sites in Yellowstone National Park have been invaded by C. arvense (Turner et al., 1997). Turner et al. (1997) reported that C. arvense was the most abundant exotic perennial at a burn site near Yellowstone Lake where the plant reached densities of approximately 1100 stems per hectare. The abundance of C. arvense increased with time in all burn severities and the density increased with fire severity (Turner et al., 1997; Crawford et al., 2001). In Mesa Verde National Park in Colorado, C. arvense aggressively invaded bare mineral soils following two major fires occurring in 1989 and 1996 (Floyd et al., 2001). Aerial seeding and herbicide treatments were utilized in an attempt to curtail the invasion of C. arvense, with the latter being the more successful (Floyd et al., 2001). Many exotic plants, including C. arvense, have invaded the National Elk Refuge in Wyoming (Matson, 2000). The primary focus of the refuge was protecting elk, which resulted in overabundance of the ungulates, rendering areas with reduced tree and shrub cover prone to exotic plant invasion (Matson, 2000).

Threatened Species

Top of page
Threatened SpeciesConservation StatusWhere ThreatenedMechanismReferencesNotes
Astragalus schmolliae (Schmoll's milkvetch)CR (IUCN red list: Critically endangered) CR (IUCN red list: Critically endangered); NatureServe NatureServe; USA ESA candidate species USA ESA candidate speciesColoradoEcosystem change / habitat alterationUS Fish and Wildlife Service, 2015a
Centrocercus minimus (Gunnison sage-grouse)USA ESA listing as threatened species USA ESA listing as threatened speciesColorado; UtahEcosystem change / habitat alterationUS Fish and Wildlife Service, 2013
Cirsium wrightii (Wright's marsh thistle)NatureServe NatureServe; USA ESA candidate species USA ESA candidate speciesArizona; New MexicoCompetition (unspecified); Ecosystem change / habitat alterationUS Fish and Wildlife Service, 2015b
Gaura neomexicana subsp. coloradensis (Colorado butterfly plant)NatureServe NatureServe; USA ESA listing as threatened species USA ESA listing as threatened speciesColorado; Nebraska; WyomingCompetition - monopolizing resourcesUS Fish and Wildlife Service, 2012a
Sidalcea nelsonianaUSA ESA listing as threatened species USA ESA listing as threatened speciesOregon; WashingtonCompetition - monopolizing resourcesUS Fish and Wildlife Service, 2012b
Silene spaldingii (Spalding's catchfly)USA ESA listing as threatened species USA ESA listing as threatened speciesIdaho; Montana; Oregon; WashingtonCompetition - monopolizing resources; Competition - smotheringUS Fish and Wildlife Service, 2007

Social Impact

Top of page C. arvense affects aesthetic and recreational values in landscape areas and backyards (Donald, 1990; Randall and Marinelli, 1996; Gao et al., 1999). It has been observed along foot trails in national parks, and in the Peace-Athabasca delta (northern Canada) the presence of C. arvense has been reported as an annoyance to hikers (Turner et al., 1997; Wein et al., 1992).

In the past, C. arvense has been used beneficially as a medicinal and edible herb (Rogers, 1928).

Risk and Impact Factors

Top of page Invasiveness
  • Invasive in its native range
  • Proved invasive outside its native range
  • Highly adaptable to different environments
  • Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
  • Highly mobile locally
  • Has high reproductive potential
  • Has propagules that can remain viable for more than one year
Impact outcomes
  • Damaged ecosystem services
  • Ecosystem change/ habitat alteration
  • Negatively impacts agriculture
  • Negatively impacts animal health
  • Negatively impacts tourism
  • Reduced amenity values
  • Reduced native biodiversity
Impact mechanisms
  • Competition - monopolizing resources
  • Competition
  • Pest and disease transmission
  • Produces spines, thorns or burrs
Likelihood of entry/control
  • Highly likely to be transported internationally accidentally
  • Difficult/costly to control

Uses List

Top of page

Materials

  • Miscellaneous materials
  • Poisonous to mammals

Medicinal, pharmaceutical

  • Traditional/folklore

Prevention and Control

Top of page Introduction

Donald (1990) summarized the management of C. arvense using non-chemical methods and herbicides in various crops. Edwards et al. (2000) provides more recent insights to promote interspecific competition to manage C. arvense through timing of crop sowing, grazing regimes, and nitrogen fertilization. Another useful review proposing integrated strategies for control of C. arvense in crops involving tillage, herbicide use and cultural control is provided by Pollack and Bailey (2001).

Regulatory Control

C. arvense has been named on most state and federal seed and weed noxious weed laws in the USA. Canadian legislation is similar (Moore, 1975). It appeared on more noxious weeds lists (33) than any other weed in North America (Skinner et al., 2000). C. arvense was introduced into French Canada from Europe (Anon., 1918) before being spread into Vermont and New York in the USA (Stevens, 1846). Detmers (1927) concluded that it must have been introduced before 1795 because a Vermont law was enacted that year to halt its spread. By 1844, Ohio law limited sale of seed contaminated with C. arvense and required landowners to mow infested land and adjacent roadsides (Detmers, 1927). Judging by its current distribution in North America, state and federal legislation has been somewhat ineffective in limiting the spread of the weed (Wilson, 1981a; Skinner et al., 2000). It is also regulated in the UK under the 1959 Weeds Act (Bond and Turner, 2003).

Cultural Control and Sanitary Methods

Combining herbicides with cultivation, mowing or grazing, and competitive crops is more effective for controlling C. arvense than herbicides alone. Various combinations have been tested and reviewed for selective control of C. arvense in the major field crops (Donald, 1990; Edwards et al., 2000). Cultural practices used alone are frequently ineffective. Even when combinations of control practices are used, repeated control measures over several years are required to reduce the severity of the problem (Donald, 1990, 1992, 1994; Donald and Prato, 1992a, 1992b). However, more recent work has identified more specific approaches that may provide effective control. Cover crops show some potential (Moyer et al., 2000). Mowing or grazing needs to be adjusted to levels appropriate for a given system (Wilson and Kachman, 1999; Eerens et al., 2002). Edwards et al. (2000) recommended sowing crops of competing species as soon as possible after cultivation. Populations of C. arvense in field margins were also reduced by sowing other species (West et al., 1997; Denys and Tschamtke, 2002). For example, Ominski et al. (1999) found that Medicago sativa effectively suppressed C. arvense, resulting in more patchy populations. Fertilization, particularly with N may also reduce C. arvense populations, particularly in the absence of grazing (Edwards et al., 2000). Repeated mowing in combination with sowing of perennial grasses has been shown to virtually eliminate C. arvense (Wilson and Kachman, 1999). Cormack (2002) achieved 75% reduction in shoots following mowing in a legume crop. In New Zealand pastures, mowing later in the season resulted in greater reduction in autumnal root biomass (Bourdot et al., 1998). Mitchell et al. (2002) observed that grazing by sheep 3-4 times approximately 3 weeks between grazing depleted root reserves of C. arvense sufficiently to prevent survival. Van Toor and Popay (1995) found that wounding C. arvense plants in advance of grazing increased grazing pressure by sheep by improving palatability. Recent advances in knowledge of the biology of C. arvense, such as the development of shoot emergence models (Donald, 2000; Jensen et al., 2002) should aid in devising management strategies.



Biological Control

Maw (1976) and Moore (1975) summarized information on insects found on shoots and roots of C. arvense. For additional information on insects and nematodes found on C. arvense, see Natural Enemies.

Survey work has identified numerous potential native biological control agents for C. arvense (Watson and Keogh, 1980; Perju et al., 1995). Biological agents for controlling C. arvense have been reviewed (Andres, 1980; Peschken et al., 1980; Trumble and Kok, 1982; Monnig, 1987). Widespread adoption of foreign biological control agents is unlikely because of public concern for native thistles (Peschken and Beecher, 1973) and the general lack of effectiveness of currently available biological control agents. Unfortunately, many of the insects and nematodes listed (see Natural Enemies) are often widespread, persistent pests of important crop species, limiting their use on commercial farms.

The weevil Ceutorhynchus litura severely reduced overwintering survival of below-ground adventitious shoots of C. arvense to as little as 3% of that of uninfested shoots in Canada (Peschken and Beecher, 1973). In a 3-year study in Montana, 8 to 12% of weevil-infested shoots survived from one year to the next compared with 94 to 99% of uninfested shoots (Rees, 1990). This stem feeder was not nearly as devastating in spring as in autumn. Weevil damage also promoted the invasion of damaged shoots by other arthropods (mites, spiders, springtails), nematodes, and fungi, although the role of these organisms in plant death was not determined. The insect may have assisted in spreading the rust fungus, Puccinia punctiformis (Peschken and Beecher, 1973), although this assertion was not substantiated later (Peschken and Wilkinson, 1981). C. litura was released 18 times in the USA in California, Colorado, Idaho, Montana, New Jersey, South Dakota, and Washington between 1971 and 1975. In Montana, C. litura spread 9 km in 10 years and the proportion of infested plants increased from 11 to 29% in 1977 to over 80% after 10 years. In Canada, this insect did not greatly or consistently increase mortality of C. arvense shoots (Peschken and Wilkinson, 1981). A weevil, Larinus planus, that feeds on seed heads of C. arvense was accidently introduced into the USA, and may be useful for controlling seed production to prevent large areas of infestation from expanding (Drlik et al., 2000). However, it has been shown to attack a native thistle, Cirsium undulatum var. tracyi in Colorado (Louda and O'Brien, 2002). Rhiocyllus conicus, a weevil that plays a similar role is no longer favoured for biological control because it also attacks native thistles (Drlik et al., 2000). Predispersal seed predation by Dasyneura gibsoni and Orellia ruficanda can significantly reduce seed output (Forsyth and Watson, 1985; Heimann and Cussans, 1996). Cassida rubiginosa is a fairly effective control agent, and may work well in combination with the effects of competition on C. arvense using plants such as Festuca arundinacea and Coronilla varia (Ang et al., 1995).

Fungi and higher plant parasites found on C. arvense have been reviewed (Moore, 1975; see also Natural Enemies). Most pathogenic viruses or bacteria reported on C. arvense are diseases of crops, such as the tobacco rattle tobravirus (Cooper and Harrison, 1973), limiting their potential for biological control of C. arvense. C. arvense is an alternative host for diseases and nematodes of crops. Drlik et al. (2000) list Pseudomonas syringae pv. tagetis, Puccinia punctiformis, and Sclerotinia scloerotiorum as the three main pathogens under investigation for use against C. arvense in North America. None of these were yet available commercially. Pseudomonas syringae pv. tagetis showed limited ability to affect Canada thistle survival in one USA study (Gronwald et al., 2002), but in Minnesota it reduced C. arvense biomass significantly in conservation tillage systems (Hoeft et al., 2001). Work in Germany has suggested that control with P. punctiformis could prevent flowering and hinder several years' growth (Kluth et al., 2003), and shows some potential for development into a mycoherbicide (Bond and Turner, 2003). The fungus Sclerotinia sclerotiorum applied to C. arvense patches as a biological control agent killed 20 to 80% of shoots in Montana (Brosten and Sands, 1986). Shoot emergence was also severely reduced in the growing season following treatment. Defoliation of shoots has a debilitating effect on the root systems as well (Bourdot and Harvey, 1994). S. sclerotiorum is an aggressive, persistent pathogen on many broadleaved crop species, limiting its use on commercial farms. A study in the Netherlands showed that the risks associated with S. sclerotinium may be manageable, however (Bourdot et al., 2001). Harvey et al. (1998) investigated using auxotrophic strains of S. sclerotinium to decrease the risk of pathogenic effects on crops, but these strains were less effective against C. arvense. Infestation of C. arvense by the bacterium P. syringae tagetis results in stunted plants that are unable to flower and are less able to overwinter (Drlik et al., 2000). Three insects that feed on C. arvense, (Aphis fabae spp. Cirsiiacanthoidis, Uroleucon cirsii and the beetle Cassida rubiginosa) were found to transmit the fungal pathogen P. punctiformis (Kluth et al., 2002), indicating the possibility of using synergism in biological control efforts. Furthermore, Bacher et al. (2002) showed that development of the beetle Apion onopordi was improved in plants infested with P. punctiformis, which is in turn is promoted by A. onopordi (Friedli and Bacher, 2001). However, Kruess (2002) found that the combination of Cassida rubiginosa and the pathogen Phoma destructive provided less efficient control of C. arvense. Green et al. (2001) observed high disease ratings for infection by Alternaria cirsinoxia in Saskatchewan, Canada.

Chemical Control

Chemical control of C. arvense has been reviewed (Moore, 1975; Donald, 1990). Different growth stages differ greatly in susceptibility to herbicides. Plants in the rosette stage are more susceptible than plants that have already bolted (Hunter, 1996; Miller and Lym, 1998); seedlings are more susceptible and sensitive to a greater variety of herbicides than mature plants (Vangessel, 1999). With the advent of herbicide-resistant crops, new possibilities for control of C. arvense within crops have appeared, including use of glyphosate, which does provide effective control in these systems (May, 2000; Sarpe et al., 2001).

Herbicides that have been used in different systems:

Temperate cereals
Bromoxynil, chlorsulfuron, clopyralid, 2,4-D, dicamba, MCPA, metsulfuron, flurasolum, Iodosulfuron-methyl-sodium
foramsulfuron.

Maize and/or sorghum
Atrazine, bentazone, bromoxynil, clopyralid, 2,4-D, dicamba.

Soyabeans
Acifluorfen, bentazone.

Sugarbeets
Clopyralid.

Dry Beans
Bentazone.

Peas
Bentazone, MCPA.

Pasture
Bromoxynil, chlorsulfuron, 2,4-D, dicamba, metsulfuron, picloram, hexazinone.

Chemical fallow
Atrazine, chlorsulfuron, 2,4-D, dicamba, glyphosate, metsulfuron, picloram.

Non-Cropland
Amitrole, atrazine, bromoxynil, chlorsulfuron, 2,4-D, dicamba, dichlobenil, glyphosate, hexazinone, imazapyr, metsulfuron, picloram, sulfometuron, tebuthiuron.

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