Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Parthenium hysterophorus
(parthenium weed)

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Datasheet

Parthenium hysterophorus (parthenium weed)

Summary

  • Last modified
  • 27 September 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Host Plant
  • Preferred Scientific Name
  • Parthenium hysterophorus
  • Preferred Common Name
  • parthenium weed
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Plantae
  •     Phylum: Spermatophyta
  •       Subphylum: Angiospermae
  •         Class: Dicotyledonae
  • Summary of Invasiveness
  • P. hysterophorus is an annual herb that aggressively colonises disturbed sites. It is considered as one of the ‘100 most invasive species in the world’ by the IUCN (G...

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Pictures

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PictureTitleCaptionCopyright
P. hysterophorus plant and flowers.
TitleWhole plant and inflorescence
CaptionP. hysterophorus plant and flowers.
Copyright©S.D. Sawant
P. hysterophorus plant and flowers.
Whole plant and inflorescenceP. hysterophorus plant and flowers.©S.D. Sawant
Parthenium weed: detail of capitula (flower).
TitleDetail of flower
CaptionParthenium weed: detail of capitula (flower).
Copyright©CABI BioScience
Parthenium weed: detail of capitula (flower).
Detail of flowerParthenium weed: detail of capitula (flower).©CABI BioScience
Parthenium weed: rosette stage.
TitleRosette stage
CaptionParthenium weed: rosette stage.
Copyright©CABI BioScience
Parthenium weed: rosette stage.
Rosette stageParthenium weed: rosette stage.©CABI BioScience
Parthenium weed: rosette stage.
TitleRosette stage
CaptionParthenium weed: rosette stage.
Copyright©CABI BioScience
Parthenium weed: rosette stage.
Rosette stageParthenium weed: rosette stage.©CABI BioScience
Parthenium weed: mature flowering stage infesting crops along the Ganges, India.
TitleCrop infestation by P. hysterophorus
CaptionParthenium weed: mature flowering stage infesting crops along the Ganges, India.
Copyright©CABI BioScience
Parthenium weed: mature flowering stage infesting crops along the Ganges, India.
Crop infestation by P. hysterophorusParthenium weed: mature flowering stage infesting crops along the Ganges, India.©CABI BioScience

Identity

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Preferred Scientific Name

  • Parthenium hysterophorus L.

Preferred Common Name

  • parthenium weed

Other Scientific Names

  • Argyrochaeta bipinnatifida Cav.
  • Argyrochaeta parviflora Cav.
  • Parthenium glomeratum Rollins
  • Parthenium lobatum Buckl.

International Common Names

  • English: barley flower; bastard feverfew; bitterweed; broomweed; carrot grass; congress grass; congress weed; dog flea weed; false ragweed; featherfew; feverfew; mugwort; Paterson’s curse; ragweed parthenium; Santa Maria feverfew; star weed; white top; white top weed; whiteheads; wild wormwood; wormwood
  • Spanish: ajenjo cimarron; amargosa; camalote; escoba amarga; hierba amargosa; istafiate; requeson
  • French: fausse camomille; matricaire; parthenium matricaire
  • Portuguese: mentruz

Local Common Names

  • Belize: coriente; siiu; silantro
  • Brazil: coentro-do-mato; fazendeiro; losna-branca
  • Caribbean: feverfew
  • China: yin jiao ju
  • Cuba: artemisilla; cofitillo; escoba amarga
  • Dominican Republic: baille lame; escoba de Puerco; escobita amarga; friega platos; yerba amarga; yerba blanca; yerba de burro; yerba mala
  • Ethiopia: arama-kuba; arama-sorgo; biyabassa; chebchabe; dayessa; faramssissa; kalignole; qinche; terekabi
  • Guatemala: hauay; tacana
  • Haiti: absinthe marron; balai amer; feuilles bauton; feuilles bouto; parthene multifide
  • Honduras: ajenjo; escobilla
  • India: bhoothkeda; carrot weed; chatak chandani; Coimbatore chedi; congree grass; congress pacha; gajar ghas; gazar ghas; keepa geda; osadi
  • Jamaica: dog-flea weed; whitetop
  • Mexico: altamisa; altamisa cimarrona; altamisa del campo; altamisilla; arrocillo; artaniza; chaile; cicutilla; cola de ardilla; confitillo; falsa altamisa; hauay; hierba amargosa; hierba del burro; hierba del gusano; huachichole; jihuite amargo; manzanilla del campo; romerillo; tzail-cuet; tzaile; yerba de la oveja; zacate amargo
  • New Caledonia: fausse camomille
  • Nicaragua: manzanilla montera
  • Pakistan: babyflower; gandhi booti; lewanai bhang
  • Puerto Rico: ajenjo cimarrón; artemisa cimarrona; yerba amarga
  • Saint Lucia: matnitjen; whitehead
  • South Africa: famine weed
  • Sweden: Flikparthenium
  • USA: false ragweed; quinine-weed; ragweed parthenium; Santa María

EPPO code

  • PTNHY (Parthenium hysterophorus)

Summary of Invasiveness

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P. hysterophorus is an annual herb that aggressively colonises disturbed sites. It is considered as one of the ‘100 most invasive species in the world’ by the IUCN (GISD, 2018). Native to the New World, it has been accidentally introduced into several countries and has become a serious agricultural and rangeland weed in parts of Australia, Asia, Africa and the Pacific Islands. It is reported as a major weed in field crops in more than forty different countries (Bajwa et al., 2016), with yield losses estimated in millions of dollars in Australia (Kaur et al., 2014). It grows on any type of soil and in a wide range of habitats. It affects the production of crops, animals, human and animal health, and biodiversity. Several characteristics, such as wide adaptability, photo- and thermo-insensitivity, lack of natural enemies in non-native regions, drought tolerance, strong competition and allelopathy, high seed production ability, longevity of seeds in soil seed banks, and small and light seeds that are capable of long distance travel via wind, water, birds, vehicles, farm machinery and other animal traffic, contribute to its rapid introduction world-wide, cutting across national boundaries and climate barriers (Kaur et al., 2014; Bajwa et al., 2016). The genetic diversity found among different populations and biotypes are also strongly contributing to its invasion success (Bajwa et al., 2018).

The species is reported as invasive in various countries in Asia, Africa, and Oceania (Gnanavel, 2013; EPPO, 2018; GISD, 2018; PIER, 2018). In the Americas, it is reported as invasive in Cuba, and in Trinidad and Tobago. In Cuba it is considered as one of the most noxious species (Oviedo Prieto et al., 2012). Although listed as introduced by various sources it is also listed as native by others (USDA-ARS, 2018). In Trinidad and Tobago it is a predominant weed of industrial areas, crops, orchards, ornamentals and greenhouses (Bridgemohan et al., 2015).

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Plantae
  •         Phylum: Spermatophyta
  •             Subphylum: Angiospermae
  •                 Class: Dicotyledonae
  •                     Order: Asterales
  •                         Family: Asteraceae
  •                             Genus: Parthenium
  •                                 Species: Parthenium hysterophorus

Notes on Taxonomy and Nomenclature

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The Asteraceae (Compositae) is the largest flowering plants family with more than 23,600 species distributed worldwide (Encyclopedia of Life, 2018). Several species are known for their economic value as ornamentals, cut-flowers, for food and beverage uses. Some species have known ethnobotanical uses.

Parthenium is a genus from the New World with ca. 16 species, including P. argentatum, used as a rubber substitute and P. hysterophorus, one of the most invasive species worldwide (Ray, 1993; Flora of North America, 2018; GISD, 2018). 

The name Parthenium hysterophorus is universally accepted for this increasingly widespread weed, commonly known as parthenium weed. P. pinnatifidum is an illegitimate name reported for P. hysterophorus. Villanova binnatifida is an unresolved name associated with a synonym of P. hysterophorus (The Plant List, 2013). There are different races and biotypes of P. hysterophorus found in several countries, and which have differing flowering patterns and reproductive behaviour. This is discussed in the Biology and Ecology section of this datasheet.

Description

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P. hysterophorus is an erect, much-branched with vigorous growth habit, aromatic, annual (or a short-lived perennial), herbaceous plant with a deep taproot. The species reproduces by seed. In its neotropical range it grows to 30-90 cm in height (Lorenzi, 1982; Kissmann and Groth, 1992), but up to 1.5 m, or even 2.5 m, in exotic situations (Haseler, 1976; Navie et al., 1996). Shortly after germination the young plant forms a basal rosette of pale green, pubescent, strongly dissected, deeply lobed leaves, 8-20 cm in length and 4-8 cm in width. The rosette stage may persist for considerable periods during unfavourable conditions (such as water or cold stress). As the stem elongates, smaller, narrower and less dissected leaves are produced alternately on the pubescent, rigid, angular, longitudinally-grooved stem, which becomes woody with age. Both leaves and stems are covered with short, soft trichomes, of which four types have been recognized and are considered to be of taxonomic importance within the genus (Kohli and Rani, 1994).

Flower heads are both terminal and axillary, pedunculate and slightly hairy, being composed of many florets formed into small white capitula, 3-5 mm in diameter. Each head consists of five fertile ray florets (sometimes six, seven or eight) and about 40 male disc florets. The first capitulum forms in the terminal leaf axil, with subsequent capitula occurring progressively down the stem on lateral branches arising from the axils of the lower leaves. Thousands of inflorescences, forming in branched clusters, may be produced at the apex of the plant during the season. Seeds (achenes) are black, flattened, about 2 mm long, each with two thin, straw-coloured, spathulate appendages (sterile florets) at the apex which act as air sacs and aid dispersal.

Plant Type

Top of page Annual
Broadleaved
Herbaceous
Seed propagated

Distribution

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There is some uncertainty about the extent of the native range of P. hysterophorus in the New World. It is regarded by Bajwa et al. (2016) as native to the Americas, introduced elsewhere. Acevedo-Rodríguez and Strong (2012) report the species as native to North America, Central America, South America and the West Indies. At the National Commission of Biodiversity of Mexico (CONABIO), the species is listed as originating in the east of Mexico and the Antilles, with a secondary native distribution from the southern USA to South America (CONABIO, 2018). Other sources report that the species originated in the region surrounding the Gulf of Mexico, including southern USA, or in central South America (Dale, 1981; Navie et al., 1996), being now widespread in North and South America and the Caribbean, and Fournet and Hammerton (1991) indicate that it occurs in 'probably all islands' of the Lesser Antilles.

Since its accidental introduction into Australia and India in the 1950s, probably as a contaminant of grain or pasture seeds, it has achieved major weed status in those countries. It was first recorded in southern Africa in 1880 but was not reported as a common weed in parts of that region until the mid-1980s following extensive flooding on the east coast (McConnachie et al., 2011). Recent reports of the weed from other countries indicate that its geographic range continues to increase.

The species is present in Asia, Africa, North America, Central America, the Caribbean, South America, Europe and Oceania (See Distribution Table for details: Acevedo-Rodríguez and Strong, 2012; EPPO, 2018; PIER, 2018; Missouri Botanical Garden, 2018; USDA-ARS, 2018). In Europe P. hysterophorus is considered as an ephemeral species (The Euro+Med Plantbase, 2018). 

Distribution Table

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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

BangladeshPresentIntroduced Invasive Mahadevappa, 1997; EPPO, 2014
BhutanPresentIntroduced Invasive Parker, 1992; EPPO, 2014
ChinaPresentIntroduced Invasive Aneja et al., 1991; EPPO, 2014; PIER, 2018
-GuangdongPresentIntroducedAneja et al., 1991; EPPO, 2014; PIER, 2018
-GuangxiPresentIntroducedAneja et al., 1991; EPPO, 2014; PIER, 2018
-GuizhouPresentIntroducedMissouri Botanical Garden, 2008; EPPO, 2014; PIER, 2018
-HunanPresentIntroducedAneja et al., 1991; EPPO, 2014
-YunnanPresentIntroducedAneja et al., 1991; EPPO, 2014; PIER, 2018
IndiaWidespreadIntroduced Invasive Holm et al., 1991; EPPO, 2014
-Andhra PradeshWidespreadIntroducedSantapau, 1967; Ellis and Swaminathan, 1969; Mahadevappa, 1997
-AssamPresentIntroducedRao, 1979; Kohli and Rani, 1994; EPPO, 2014
-BiharWidespreadIntroducedChandra, 1973; Maheshwari and Pandey, 1973; EPPO, 2014
-ChandigarhWidespreadIntroducedKumari et al., 1985; Aneja et al., 1991; EPPO, 2014
-DelhiWidespreadIntroducedMaheshwari, 1966; Kohli and Rani, 1994; EPPO, 2014
-GujaratWidespreadIntroducedMahadevappa, 1997; EPPO, 2014
-HaryanaWidespreadIntroducedAneja et al., 1991; EPPO, 2014
-Himachal PradeshWidespreadIntroducedVaid and Naithani, 1970; EPPO, 2014
-Indian PunjabWidespreadIntroducedMahadevappa, 1997; EPPO, 2014
-Jammu and KashmirWidespreadIntroducedHakoo, 1963; Mahadevappa, 1997; EPPO, 2014
-KarnatakaWidespreadIntroducedJayachandra, 1971; Mahadevappa, 1997; EPPO, 2014
-KeralaWidespreadIntroducedMahadevappa, 1997; EPPO, 2014
-Madhya PradeshWidespreadIntroducedMaheshwari, 1968; Tiwari and Bisen, 1984; EPPO, 2014
-MaharashtraWidespreadIntroducedRao, 1956; Vartak, 1968; EPPO, 2014
-OdishaPresentIntroducedMahadevappa, 1997; EPPO, 2014
-RajasthanPresentIntroducedGena and Bhardwaj, 1980; Mahadevappa, 1997; EPPO, 2014
-Tamil NaduWidespreadIntroducedEllis and Swaminathan, 1969; Mahadevappa, 1997; EPPO, 2014
-Uttar PradeshWidespreadIntroducedEllis and Swaminathan, 1969; Mahadevappa, 1997; EPPO, 2014
-West BengalWidespreadIntroducedMandal et al., 1980; Mahadevappa, 1997; EPPO, 2014
IsraelWidespreadIntroduced Invasive Joel and Liston, 1986; Navie et al., 1996; EPPO, 2014; Euro+Med PlantBase, 2018
JapanPresentEPPO, 2014; PIER, 2018
-Ryukyu ArchipelagoPresentIntroducedUSDA-ARS, 2012
JordanPresent, few occurrencesIntroduced Not invasive Euro+Med PlantBase, 2018
Korea, Republic ofPresent Invasive Shabbir and Adkins, 2013; EPPO, 2014
MalaysiaPresentIntroduced Invasive Rezaul Karim, 2014
NepalPresentIntroduced Invasive Evans, 1997a; Aneja et al., 1991; Mishra, 1991; EPPO, 2014; India Biodiversity Portal, 2018
OmanPresentIntroduced1998 Invasive Alhammadi, 2010; EPPO, 2014
PakistanPresentIntroduced1980s Invasive Shabbir et al., 2011; EPPO, 2014
Sri LankaPresentJayasurya, 2005; Kelaniyangoda and Ekanayake, 2008; EPPO, 2014
TaiwanPresentIntroducedTowers and Mitchell, 1983; Peng et al., 1988; Navie et al., 1996; EPPO, 2014
VietnamPresentIntroduced Invasive Maheshwari and Pandey, 1973; Aneja et al., 1991; Navie et al., 1996; EPPO, 2014
YemenPresentIntroduced Invasive Alhammadi, 2010; EPPO, 2014

Africa

ComorosPresentIntroducedMissouri Botanical Garden, 2008; EPPO, 2014; India Biodiversity Portal, 2018
EgyptPresentIntroduced Invasive Zahran and Willis, 2009; EPPO, 2014
EritreaPresentIntroducedUSDA-ARS, 2012; EPPO, 2014
EthiopiaWidespreadIntroduced1980s Invasive Evans, 1997a; Medhin, 1992; Fasil, 1994; Frew et al., 1996; EPPO, 2014
KenyaPresentIntroduced Invasive Ivens, 1989; Njoroge, 1989; Navie et al., 1996; EPPO, 2014
MadagascarPresentIntroduced Invasive Aneja et al., 1991; EPPO, 2014; India Biodiversity Portal, 2018
MauritiusWidespreadIntroduced Invasive Holm et al., 1991; Navie et al., 1996; Mahadevappa, 1997; EPPO, 2014; PIER, 2018
MayottePresentIntroducedUSDA-ARS, 2012; EPPO, 2014
MozambiquePresentIntroduced Invasive Aneja et al., 1991; EPPO, 2014
RéunionPresentIntroduced Invasive Navie et al., 1996; Mahadevappa, 1997; EPPO, 2014
SeychellesPresentIntroduced Invasive Navie et al., 1996; Mahadevappa, 1997; EPPO, 2014
SomaliaPresentIntroducedTamado and Milberg, 2000; EPPO, 2014
South AfricaPresentIntroduced Invasive Maheshwari, 1966; Picman and Towers, 1982; Navie et al., 1996; McConnachie et al., 2011; EPPO, 2014; USDA-ARS, 2018
SwazilandPresentIntroducedHenderson, 2001; EPPO, 2014
TanzaniaPresentIntroduced Invasive McConnachie et al., 2011; EPPO, 2014
UgandaPresentIntroduced Invasive McConnachie et al., 2011; EPPO, 2014
ZimbabwePresentIntroducedMcConnachie et al., 2011; EPPO, 2014

North America

BermudaPresentNative Not invasive Dale, 1981; Aneja et al., 1991; EPPO, 2014
MexicoWidespreadNative Not invasive Haseler, 1976; Dale, 1981; Aneja et al., 1991; Holm et al., 1991; EPPO, 2014
USAWidespreadNative Not invasive Dale, 1981; Holm et al., 1991; EPPO, 2014; USDA-ARS, 2018
-AlabamaPresentAneja et al., 1991; Kohli and Rani, 1994; EPPO, 2014
-ArkansasPresentEPPO, 2014
-ConnecticutPresentEPPO, 2014
-DelawarePresentIntroducedUSDA-ARS, 2012; EPPO, 2014
-District of ColumbiaPresentIntroducedUSDA-ARS, 2012; EPPO, 2014
-FloridaPresentAneja et al., 1991; Kohli and Rani, 1994; EPPO, 2014
-HawaiiPresentIntroduced Invasive PIER, 2008; USDA-ARS, 2012; EPPO, 2014Hawai’i, Jaua ‘i, Maui, Moloka’i, O’ahu
-IllinoisPresentFernold, 1970; Mahadevappa, 1997; EPPO, 2014
-KansasPresentFernold, 1970; Mahadevappa, 1997; EPPO, 2014
-LouisianaPresentMahadevappa, 1997; EPPO, 2014
-MarylandPresentArny, 1897; Kohli and Rani, 1994; EPPO, 2014
-MassachusettsPresentArny, 1897; EPPO, 2014
-MichiganPresentFernold, 1970; Mahadevappa, 1997; EPPO, 2014
-MinnesotaPresentMackoff and Dahl, 1951; Mahadevappa, 1997
-MississippiPresentIntroducedUSDA-ARS, 2012; EPPO, 2014
-MissouriPresentFernold, 1970; Mahadevappa, 1997; EPPO, 2014
-New JerseyPresentIntroducedUSDA-ARS, 2012; EPPO, 2014
-New MexicoPresentIntroducedUSDA-ARS, 2012
-New YorkPresentEPPO, 2014
-OhioPresentFernold, 1970; Mahadevappa, 1997; EPPO, 2014
-OklahomaPresentEPPO, 2014
-PennsylvaniaPresentEPPO, 2014
-South CarolinaPresentEPPO, 2014
-TexasPresentCastex et al., 1940; McClay et al., 1995; Mahadevappa, 1997; EPPO, 2014
-VirginiaPresentArny, 1897; Mahadevappa, 1997; EPPO, 2014

Central America and Caribbean

AnguillaPresentNativeUSDA-ARS, 2008; EPPO, 2014
Antigua and BarbudaPresentNativeUSDA-ARS, 2012; EPPO, 2014
ArubaPresentNativeUSDA-ARS, 2012; EPPO, 2014
BahamasPresentNativeUSDA-ARS, 2012; EPPO, 2014
BarbadosPresentNativeDale, 1981; Kohli and Rani, 1994; USDA-ARS, 2012; EPPO, 2014
BelizePresentNative Not invasive Dale, 1981; Aneja et al., 1991; EPPO, 2014; Missouri Botanical Garden, 2018
British Virgin IslandsPresentNativeAcevedo-Rodríguez and Strong, 2012; Missouri Botanical Garden, 2018Tortola, Virgin Gorda
Cayman IslandsPresentNativeUSDA-ARS, 2012; EPPO, 2014
Costa RicaPresentNative Not invasive Dale, 1981; Parsons and Cuthbertson, 1992; EPPO, 2014
CubaWidespreadIntroduced Invasive Evans, 1997b; Holm et al., 1991; Navie et al., 1996; Oviedo Prieto et al., 2012; EPPO, 2014; Missouri Botanical Garden, 2018; USDA-ARS, 2018
CuraçaoPresentNative Not invasive Dale, 1981; Kohli and Rani, 1994
DominicaPresentNative Not invasive Dale, 1981; Kohli and Rani, 1994; EPPO, 2014
Dominican RepublicWidespreadNative Not invasive Evans, 1997b; Ciferri, 1956; Dale, 1981; Holm et al., 1991; EPPO, 2014
GrenadaPresentNativeUSDA-ARS, 2012; EPPO, 2014
GuadeloupePresentNative Not invasive Dale, 1981; Kohli and Rani, 1994; EPPO, 2014
GuatemalaPresentNative Not invasive Dale, 1981; Aneja et al., 1991; Kohli and Rani, 1994; EPPO, 2014; Missouri Botanical Garden, 2018
HaitiPresentNative Not invasive Dale, 1981; Aneja et al., 1991; EPPO, 2014
HondurasPresentNative Not invasive Dale, 1981; Aneja et al., 1991; Kohli and Rani, 1994; EPPO, 2014; Missouri Botanical Garden, 2018
JamaicaWidespreadNative Not invasive Dale, 1981; Aneja et al., 1991; Holm et al., 1991; Mahadevappa, 1997; EPPO, 2014
MartiniquePresentNative Not invasive Dale, 1981; Kohli and Rani, 1994; EPPO, 2014
MontserratPresentNativeUSDA-ARS, 2018
Netherlands AntillesPresentNativeUSDA-ARS, 2012; EPPO, 2014
NicaraguaPresentLewis et al., 1988; EPPO, 2014; Missouri Botanical Garden, 2018
PanamaPresentHammel, 1997; EPPO, 2014
Puerto RicoWidespreadNative Not invasive Dale, 1981; Aneja et al., 1991; Holm et al., 1991; EPPO, 2014
SabaPresentNativeAcevedo-Rodríguez and Strong, 2012
Saint Kitts and NevisPresentNativeUSDA-ARS, 2012
Saint LuciaPresentNativeUSDA-ARS, 2012; EPPO, 2014
Saint Vincent and the GrenadinesPresentNativeUSDA-ARS, 2012
Sint EustatiusPresentNativeAcevedo-Rodríguez and Strong, 2012
Sint MaartenPresentNativeAcevedo-Rodríguez and Strong, 2012
Trinidad and TobagoWidespreadNative Not invasive Dale, 1981; Aneja et al., 1991; Holm et al., 1991; Mahadevappa, 1997; EPPO, 2014; Bridgemohan et al., 2015
Turks and Caicos IslandsPresentIntroducedGBIF, 2008; PROTA, 2018
United States Virgin IslandsPresentNativeAcevedo-Rodríguez and Strong, 2012; USDA-ARS, 2012; EPPO, 2014

South America

ArgentinaWidespreadNative Not invasive Castex et al., 1940; Dale, 1981; Aneja et al., 1991; Holm et al., 1991; EPPO, 2014
BoliviaPresentNative Not invasive Dale, 1981; Aneja et al., 1991; Kohli and Rani, 1994; EPPO, 2014
BrazilPresentNative Not invasive Dale, 1981; EPPO, 2014; Flora do Brasil, 2018
-AlagoasPresentIntroducedFlora do Brasil, 2018
-BahiaPresentIntroducedFlora do Brasil, 2018
-GoiasPresentLorenzi, 1982; EPPO, 2014; Flora do Brasil, 2018
-Mato GrossoPresentIntroducedFlora do Brasil, 2018
-Mato Grosso do SulPresentLorenzi, 1982; EPPO, 2014; Flora do Brasil, 2018
-Minas GeraisPresentLorenzi, 1982; EPPO, 2014; Flora do Brasil, 2018
-ParanaWidespreadLorenzi, 1982; Kissmann and Groth, 1992; EPPO, 2014; Flora do Brasil, 2018
-PernambucoPresentIntroducedFlora do Brasil, 2018
-Rio de JaneiroPresentLorenzi, 1982; EPPO, 2014; Flora do Brasil, 2018
-Santa CatarinaPresentLorenzi, 1982; EPPO, 2014
-Sao PauloWidespreadLorenzi, 1982; Kissmann and Groth, 1992; EPPO, 2014; Flora do Brasil, 2018
-TocantinsPresentIntroducedFlora do Brasil, 2018
ChilePresentDale, 1981; EPPO, 2014
ColombiaPresentNativeMissouri Botanical Garden, 2018Valle del Cauca
EcuadorPresentNativeMissouri Botanical Garden, 2008; USDA-ARS, 2012; EPPO, 2014; Missouri Botanical Garden, 2018
French GuianaPresentNativeUSDA-ARS, 2012; EPPO, 2014
GuyanaPresentNative Not invasive Dale, 1981; Aneja et al., 1991; Mahadevappa, 1997; EPPO, 2014
ParaguayPresentNative Not invasive Dale, 1981; Aneja et al., 1991; EPPO, 2014
PeruPresentDale, 1981; EPPO, 2014; Missouri Botanical Garden, 2018
SurinamePresentNativeUSDA-ARS, 2012; EPPO, 2014
UruguayPresentNative Not invasive Dale, 1981; Aneja et al., 1991; EPPO, 2014
VenezuelaWidespreadNative Not invasive Dale, 1981; Aneja et al., 1991; Holm et al., 1991; EPPO, 2014

Europe

BelgiumTransient: actionable, under eradicationUSDA-ARS, 2012; EPPO, 2014; Euro+Med PlantBase, 2018
PolandTransient: actionable, under eradicationEPPO, 2014; Euro+Med PlantBase, 2018

Oceania

AustraliaPresentIntroduced Invasive Dale, 1981; Holm et al., 1991; EPPO, 2014
-Australian Northern TerritoryPresentIntroduced Invasive Auld and Medd, 1987; Navie et al., 1996; EPPO, 2014
-New South WalesPresentIntroducedAuld and Medd, 1987; Navie et al., 1996; EPPO, 2014
-QueenslandWidespreadIntroduced Invasive Haseler, 1976; Navie et al., 1996; EPPO, 2014
-Western AustraliaPresentEPPO, 2014
French PolynesiaPresentIntroduced Invasive Aneja et al., 1991; EPPO, 2014; PIER, 2018Raiatea
New CaledoniaPresentIntroduced Invasive Aneja et al., 1991; PIER, 2008; EPPO, 2014Iles Loyaute, Ile Mare, Ile Oyuvea, Ile Tiga, Ile Walpole, Ile Grande Terre, Ile des Pins
Papua New GuineaTransient: actionable, under eradicationEPPO, 2014
VanuatuWidespreadIntroduced Invasive Aneja et al., 1991; Holm et al., 1991; Navie et al., 1996; EPPO, 2014; PIER, 2018

History of Introduction and Spread

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P. hysterophorus is presumed to have entered India along with food grains imported from the USA (Vartak, 1968). It was first reported in India in 1810 in Arunachal Pradesh (Gnanavel, 2013). It was later identified and described by Rao (1956) in Pune (formally Poona), in the Maharashtra district, in 1955 (Vartak, 1968), and has since spread to most of the sub-continent (Nath, 1988). At present it is reported from all the states of India (Gnanavel, 2013). It is thought to have entered Pakistan, Nepal and Bangladesh via road connections, where thousands of vehicles cross between India and these countries every day at several places. In Pakistan, P. hysterophorus was first reported from Gujarat district of the Punjab Province in 1980s (Razaq et al., 1994) and since then it has rapidly spread throughout the Province of Punjab, Islamabad Capital Territory (ICT) and parts of Khyber Pukhtunkhwa Province. The weed is thought to have entered Nepal from India (Shrestha et al., 2015) and is currently found throughout most of the lowland Tarai region that borders India, and within most of the cities and urban areas in the Dun Valleys of the Siwalik range and the Mid-hill region. It was recorded in Nepal in 1967 and as present in New Caledonia since the late 19th century (India Biodiversity Portal, 2018). It was first recorded in Sri Lanka in 1999 in the Vavuniya District.

In Australia, P. hysterophorus was first recorded near Toogoolawah in South-East Queensland  around the mid-1940’s. This population is known as the Toogoolawah type and has not spread more than 10 km (Bajwa et al., 2018). It has been suggested that this introduction was due to the movement of aircraft and machinery parts into Australia during the Second World War (Auld et al., 1983). A second accidental introduction occurred in central Queensland, north of Clermont, in 1958 and originated from contaminated pasture seed (Navie et al., 1996). This population is known as the Clermont type, which started to spread rapidly by the 1970’s. By the early 2000’s the species was covering more than 520,522 km2 (McFadyen, 1992; Bajwa et al., 2018). Both biotypes were introduced independently from the northern and southern Texas-USA races.

P. hysterophorus was first recorded from southern Africa in 1880 (McConnachie et al., 2011) but did not become a weed there until the mid-1980s. The first record from east Africa is from Ruiru, Kenya, in 1973 (PROTA, 2018), and has become a weed of Kenyan coffee plantations (Njoroge, 1991). In Ethiopia, parthenium weed was reported at Dire Dawa in 1988 (Fasil, 1994). There are two speculations: either the weed was introduced through wheat seed donated for relief from abroad or that it was introduced during the Ethio-Somali war in 1976/77 (Tamado and Milberg, 2000). However, parthenium weed was recorded in 1968 at the Alemaya University of Agriculture, Ethiopia (Tamado, 2001). From the presence of parthenium weed in Kenya and Somalia (Njoroge, 1991; Frew et al., 1996), and the capacity of parthenium weed seed to travel long distances through wind, water and other means, it is possible that it might have been introduced to Ethiopia from these neighbouring countries (Taye, 2002). According to Joshi et al. (2016) the species has spread to Kenya, Mozambique and South Africa from Ethiopia. Although P. hysterophorus is cited as introduced in Cuba by various sources (Evans, 1997b; Holm et al., 1991; Navie et al., 1996; Oviedo Prieto et al., 2012; EPPO, 2018), it is also cited as a native species by USDA-ARS (2018).

Introductions

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Introduced toIntroduced fromYearReasonIntroduced byEstablished in wild throughReferencesNotes
Natural reproductionContinuous restocking
Australia 1950s Yes No Auld et al. (1983)
Ethiopia 1960s No No Tamado (2001)
India USA 1950s Yes No Vartak (1968)
Kenya 1970s Yes No
Queensland 1958 Seed trade (pathway cause) Yes No Navie et al. (1996)
South Africa 1880 Yes No McConnachie et al. (2011)
Pakistan 1980s Hitchhiker (pathway cause) Yes No Razaq et al. (1994)
Nepal 1967 Hitchhiker (pathway cause) Yes No India Biodiversity Portal (2018)
Sri Lanka 1999 Yes No Kelaniyangod and Ekanayake (2008)

Risk of Introduction

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P. hysterophorus is an annual herb that has invaded large areas in different types of habitats and environmental conditions, mainly in Australia, Asia, Africa and the Pacific Islands (Bajwa et al., 2016). From the experience in India, Australia and Africa, there is a considerable risk of accidental introduction of the species (Bajwa et al., 2016; EPPO, 2018; GISD, 2018). It can be spread via flowing water or can be blown by wind, making prevention of spread difficult. Once introduced, it can be spread by vehicles and farm machinery, and the transport of goods, sand, soil and compost from infested areas to uninfested areas. The rapid colonisation is aided by the mobility of the seed, the adaptability of the species to a wide range of habitats, its drought tolerance, the high growth rate and the allelopathic effects it has over other species (Rubaba et al., 2017). In spite of intensified plant quarantine regulations in most countries, the high risks of introduction will persist, and given the wide climatic adaptability of the weed, further territories are likely to be affected (see, for example, McConnachie et al. (2011) for areas at risk in southern and eastern Africa).

Using CLIMEX modelling, Shabbir (2012) predicted that P. hysterophorus is likely to expand its geographical distribution range in Australia and southern Asia, particularly into southern Pakistan where additional moisture is available in the form of irrigation. The current climates of northern Africa and south and south-eastern Europe are also suitable for parthenium weed. Climate changes may alter rainfall patterns, cause rising temperatures and atmospheric carbon dioxide concentrations, all which can promote the expansion of the species range (Bajwa et al., 2016). Under climate change, the northern parts of the African continent, northern China, most of eastern and northern Europe and the Mediterranean are also under the threat of invasion. Currently, P. hysterophorus is still an infrequent weed in Europe, unlikely to become established in temperate zones.

Habitat

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P. hysterophorus is an annual herb that is especially prolific in disturbed habitats, such as along roadsides and railway tracks, river and creek banks, in stock yards, around buildings and on wasteland, from where it spreads and invades agricultural systems (EPPO, 2018; PROTA, 2018). It is also present in rangelands, coastal dunes, villages, gardens, along streams, rivers, plant nurseries and crop fields. Rashmi et al. (1999) reported that it is also present in wetlands. As reported by Kaur et al. (2014), drought and the reduced pasture cover that follows creates the ideal conditions for the species to become established. It grows from hot arid and semi-arid to humid habitats, in altitudes from sea level to 4286 metres (Rubaba et al., 2017; PROTA, 2018).

In Taiwan is widely naturalised in open areas in seashores and disturbed sites. In Mexico it is found in roadsides, pastures, near railways, occasionally in cultivated sites, and in deciduous and semideciduous forests up to 2500 m elevation (CONABIO, 2018). In India it is found in arable land, permanent crops and orchards, pastures, continental water banks, riverbanks and canal sides, roadsides, wastelands and railways (Gnanavel, 2013).

Habitat List

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CategoryHabitatPresenceStatus
Littoral
Coastal areas Present, no further details Harmful (pest or invasive)
Coastal areas Present, no further details Natural
Coastal dunes Present, no further details Harmful (pest or invasive)
Coastal dunes Present, no further details Natural
Terrestrial-managed
Cultivated / agricultural land Principal habitat Harmful (pest or invasive)
Disturbed areas Principal habitat Harmful (pest or invasive)
Disturbed areas Principal habitat Natural
Industrial / intensive livestock production systems Secondary/tolerated habitat Harmful (pest or invasive)
Managed forests, plantations and orchards Principal habitat Harmful (pest or invasive)
Managed grasslands (grazing systems) Secondary/tolerated habitat Harmful (pest or invasive)
Protected agriculture (e.g. glasshouse production) Present, no further details Harmful (pest or invasive)
Rail / roadsides Principal habitat Harmful (pest or invasive)
Rail / roadsides Principal habitat Natural
Urban / peri-urban areas Principal habitat Harmful (pest or invasive)
Urban / peri-urban areas Principal habitat Natural
Urban / peri-urban areas Principal habitat Productive/non-natural
Terrestrial-natural/semi-natural
Natural forests Present, no further details Harmful (pest or invasive)
Natural forests Present, no further details Natural
Natural grasslands Principal habitat Harmful (pest or invasive)
Natural grasslands Principal habitat Natural
Riverbanks Secondary/tolerated habitat Harmful (pest or invasive)
Riverbanks Secondary/tolerated habitat Natural
Wetlands Secondary/tolerated habitat Harmful (pest or invasive)
Wetlands Secondary/tolerated habitat Natural

Hosts/Species Affected

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P. hysterophorus is known to reduce the yield of various crops and to compete with pasture species in various countries. However, the yield loss and specific effects on the crops have not been quantified in all countries (Rubaba et al., 2017).

In Australia, the main impact of P. hysterophorus has been in the pastoral region of Queensland, where it replaces forage plants, thereby reducing the carrying capacity for grazing animals (Haseler, 1976; Chippendale and Panetta, 1994). Serious encroachment and replacement of pasture grasses has also been reported in India (Jayachandra, 1971) and in Ethiopia (Tamado, 2001Taye, 2002). The weed is also able to invade natural ecosystems, and has caused total habitat changes in native Australian grasslands and open woodlands (McFadyen, 1992).

In India, the yield losses are reported as up to 40% in several crops and a 90% reduction of forage production (Gnanavel, 2013). P. hysterophorus is now being reported from India as a serious problem in cotton, groundnuts, potatoes and sorghum, as well as in more traditional crops such as okra (Abelmoschus esculentus), brinjal (Solanum melongena), chickpea and sesame (Kohli and Rani, 1994), and is also proving to be problematic in a range of orchard crops, including vineyards, olives, cashew, coconut, guava, mango and papaya (Tripathi et al., 1991; Mahadevappa, 1997; Gnanavel, 2013).

Similar infestations of sugarcane and sunflower plantations have recently been noted in Australia (Parsons and Cuthbertson, 1992; Navie et al., 1996), whilst in Brazil and Kenya, the principal crop affected is coffee (Njoroge, 1989; Kissmann and Groth, 1992). In Ethiopia, parthenium weed was observed to grow in maize, sorghum, cotton, finger millet (Eleusine coracana), haricot bean (Phaseolus vulgaris), tef (Eragrostis tef), vegetables (potato, tomato, onion, carrot) and fruit orchards (citrus, mango, papaya and banana) (Taye, 2002). In Pakistan, the weed has been reported from number of crops, including wheat, rice, sugarcane, sorghum, maize, squash, gourd and water melon (Shabbir 2006; Shabbir et al. 2011; Anwar et al. 2012).

In Mexico, the species is reported as a weed in cotton, rice, sugarcane, Citrus spp, beans, safflower, sunflower, lentils, corn, mango, okra, bananas, tomato, grapes, alfalfa, chili peppers, luffa, marigolds and other vegetables and fruit orchards. It is also a weed in nurseries. In Argentina is reported as a weed of tobacco fields (CONABIO, 2018).

Gnanavel (2013) also reports the following detrimental effects of P. hysterophorus on crops: it inhibits nitrogen fixing bacteria in legumes; the vast quantity of pollen it produces (ca. 624 million/plants) inhibits fruit setting; it is an alternative host for viruses that cause diseases in crop plants; and it is an alternative host for mealy bugs.

Biology and Ecology

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Genetics

The chromosome number for P. hysterophorus has been reported as 2n=18 in India (Hakoo, 1963) and Australia (Navie et al., 1996); however, both lower (2n=9) and higher (2n=34) chromosome numbers have also been attributed to parthenium weed (Kohli and Rani, 1994). Germplasm collections are available at various institutions (Kew Royal Botanic Gardens, 2018; USDA-ARS, 2018). DNA information is available at the Barcode of Life Data System (BOLDS, 2018).

Two distinct races of P. hysterophorus have been identified, the 'South American' and the 'North American' races (Dale, 1981), the former with cream to yellow flowers and the latter with white flowers. Moreover, Towers et al. (1977) indicated that the sesquiterpene lactone, hymenin, which is present in plants from Argentina and Bolivia, is different from the lactone, parthenin, identified from most of the samples collected in India, as well as North and Central America. The detailed genetic comparison between plants from across the introduced range is necessary to ascertain the history of its spread and to develop management programmes.

The species has two biotypes in Australia, which are distinctive based on their introduction, the colonization ability, and their demographic spread (Adkins et al., 1997; Bajwa et al., 2018). The Toogoolawah biotype is confined to less than 10 kilometers from its point of origin. According to Hanif et al. (2011) the Toogoolawah type has a tendency to self-pollination. The Clermont biotype is invasive and has spread into more than 520,522 km2. The two biotypes also have significant genetic and morphological differences. The Clermont biotype has a higher germination rate across different environmental conditions compared with the Toogoolawah biotype, which might be contributing to its high invasive ability (Bajwa et al., 2018). Parker (1989) identified two biotypes with different flowering patterns in Mexico.

Reproductive Biology

P. hysterophorus reproduces by seeds. It is unable to reproduce vegetatively from plant parts or by apomixis, but is a prolific seed producer (15,000-25,000 achenes per plant on average, and up to 100,000 in large plants) (Haseler, 1976; Navie et al., 1996; Mahadevappa, 1997; Gnanavel, 2013), and continues to flower and fruit until senescence. Pollen grains are spheroidal, 15 to 20 μm in size, and have short to medium length spines with an average of 168,192 pollen grains produced in each capitulum (Lewis et al., 1988), adding up to about 624 million per plant (Gnanavel, 2013).

Most seeds germinate within two years if the conditions are suitable (Gnanavel, 2013). The longevity of surface-lying seeds seems to be short with little or no dormancy, but there is evidence that buried achenes can remain viable for at least 4-6 years (Navie et al., 1996). Navie et al. (1998) estimated the half-life of buried seed to be about 6 years. Gnanavel (2013) reports a longer viability for buried seeds, being able to germinate after 8-10 years. Tamado et al. (2002) report that the viability of the seeds was greater than 50% after 26 months of burial in the soil, indicating the potential build-up of a substantial and persistent soil seed bank. Germination occurrs at the mean minimum (10°C) and maximum (25°C) temperatures, as well as over a wide range of fluctuating temperatures (12/2°C - 35/25°C) in light (Tamado et al., 2002). An optimum germination temperature of 22-25°C is reported by Gnanavel (2013). It appears that the species can germinate without light, but a 12 hours photoperiod is ideal for its maximum germination (Bajwa et al., 2018). Seed germination can occur over a wide range of soil pH (2.5-10), with an optimum of 5.5-7.0 (Parsons and Cuthbertson, 1992). It can also germinate under high osmotic and salt stress (Bajwa et al., 2018). Germination may be increased after cold stratification, and with exposure to light (Karlsson et al., 2008). Flowering may begin as early as 4 weeks after seedling emergence (Jayachandra, 1971), and plants continue to flower for extended periods (6-8 months) when conditions permit. Under favourable conditions of adequate moisture and bare soil, four or five generations per year can be completed (Gnanavel, 2013).

Physiology and Phenology

P. hysterophorus is an aggressive colonizer of disturbed ground, able to germinate, grow and flower over a wide range of temperatures and photoperiods. Seeds germinate all year-round provided moisture is available and germination rate is extremely high. Four or more successive cohorts of seedlings may be produced in a season (Pandey and Dubey, 1989).

In Australia, parthenium weed germinates mainly in spring and early summer. It produces flowers and seeds throughout its life and dies in late autumn (Navie et al., 1996). It can grow at any time of the year as long as there is moisture (Tamado, 2001; Taye, 2002).

Plants emerging during the first (spring) rains usually attain a greater size and have a significantly longer lifespan than those produced in the summer. Soil moisture appears to be the major contributing factor to both the lifespan and to duration of flowering. Plant biomass production increases with increasing temperature up to an optimum day/night temperature regime of 33/22°C (Williams and Groves, 1980). Under unfavourable (dry) conditions, the life cycle may take up to 335 days, compared to 86 days under optimum conditions. Physiological studies have shown that parthenium weed has a low photorespiratory activity and has the Cphotosynthetic pathway but with positive C4 tendencies (Patil and Hegde, 1983).

The species also produces chemical compounds that when released in the soil have allelopathic effects on other species (Rubaba et al., 2017). 

Associations

In Mexico P. hysterophorus is reported as being associated with Bursera and arborescent Ipomoea species (CONABIO, 2018).

Environmental Requirements

P. hysterophorus occurs in the humid and subhumid tropics, typically favouring heavier fertile soils, such as black, alkaline clay loams, but is able to grow on a wide variety of soil types from sea level up to 2400 m (Evans, 1987a; Taye et al., 2002). Areas receiving less than 500 mm of rainfall annually are probably unsuitable, although the weed has strong adaptive methods to tolerate both moisture stress (Kohli and Rani, 1994) and saline conditions (Hegde and Patil, 1982; Khurshid et al. 2012).

Parthenium weed is an aggressive colonizer of disturbed ground, able to germinate, grow and flower over a wide range of temperatures and photoperiods. It occurs in the humid and sub-humid tropics, showing a marked preference for black, alkaline, cracking, clay soils of high fertility, but is able to grow on wide variety of soil types from sea level up to 1800 m (Evans, 1987a). In Ethiopia, it grows from low to high-mid-altitude areas at 900-2500 m asl (Taye, 2002). High clay content in soils prolonged the rosette stage, enhanced relative growth rates in height and diameter, and hampered root growth, but promoted biomass allocation to shoots (Annapurna and Singh, 2003). Mahadevappa (1997) noted that parthenium weed has several built-in properties and efficient behavioural mechanisms that enable it to overcome many ecological adversities and thus continue to survive under stress. The species is not frost tolerant, which is considered as a limiting factor to its distribution in temperate areas (Gnanavel, 2013). It grows best in subtropical regions with mean annual temperatures ranging from 10-25°C and an annual rainfall exceeding 500 mm. Areas receiving less than 500 mm of rainfall are probably unsuitable, although the weed has strong adaptive methods to tolerate both moisture stress (Kohli and Rani, 1994) and saline conditions (Hegde and Patil, 1982). The weed finds access to any type of land but it is especially prolific in disturbed habitats, for example, roadsides and railway tracks, stock-yards, around buildings and on waste land, from where it spreads and invades agricultural systems.

According to Dale (1981), the distribution of parthenium weed in Queensland is controlled by factors similar to those limiting the plant in its areas of origin. However, because of differences in land management, soils and climate, the plant covers much greater areas and is a far more significant problem in Australia. The combination of neutral to alkaline clays and absence of competing vegetation provided ideal conditions for the development of large stands of parthenium weed. The plant is more common on roadsides in Queensland than in North America, but this can be attributed to more regular disturbance. In natural grasslands, the situation is similar in the two areas, with parthenium weed becoming dominant only in the most overgrazed situations. The plant appeared more prominent in cultivated areas in North America, particularly during fallow periods, but in neither area does it significantly affect crop production. In all regions, the densest stands produce a complete ground cover with no other species present.

Climate

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ClimateStatusDescriptionRemark
As - Tropical savanna climate with dry summer Tolerated < 60mm precipitation driest month (in summer) and < (100 - [total annual precipitation{mm}/25])
Aw - Tropical wet and dry savanna climate Preferred < 60mm precipitation driest month (in winter) and < (100 - [total annual precipitation{mm}/25])
BS - Steppe climate Tolerated > 430mm and < 860mm annual precipitation
Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year
Cs - Warm temperate climate with dry summer Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers
Cw - Warm temperate climate with dry winter Preferred Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)
Af - Tropical rainforest climate Tolerated > 60mm precipitation per month
Am - Tropical monsoon climate Preferred Tropical monsoon climate ( < 60mm precipitation driest month but > (100 - [total annual precipitation(mm}/25]))

Latitude/Altitude Ranges

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Latitude North (°N)Latitude South (°S)Altitude Lower (m)Altitude Upper (m)
45 35

Air Temperature

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Parameter Lower limit Upper limit
Absolute minimum temperature (ºC) -5
Mean annual temperature (ºC) 12 25
Mean maximum temperature of hottest month (ºC) 30 40
Mean minimum temperature of coldest month (ºC) 2 12

Rainfall

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ParameterLower limitUpper limitDescription
Dry season duration06number of consecutive months with <40 mm rainfall
Mean annual rainfall5002400mm; lower/upper limits

Rainfall Regime

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Summer
Uniform

Soil Tolerances

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Soil drainage

  • free
  • impeded
  • seasonally waterlogged

Soil reaction

  • acid
  • alkaline
  • neutral

Soil texture

  • heavy
  • light
  • medium

Special soil tolerances

  • infertile
  • saline
  • shallow
  • sodic

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Aphis fabae Herbivore not specific
Bucculatrix parthenica Herbivore Leaves/Stems to genus Australia
Carmenta ithacae Herbivore Roots/Stems not specific Australia
Cercospora parthenii Pathogen Leaves not specific
Cercospora partheniphila Pathogen Leaves not specific
Conotrachelus albocinereus Herbivore Stems not specific Dhileepan and McFadyen, 2012 Australia
Entyloma compositarum Pathogen Leaves not specific
Epiblema strenuana Herbivore Leaves not specific Dhileepan and McFadyen, 2012 Australia (Queensland)
Golovinomyces cichoracearum Pathogen Leaves not specific
Insignorthezia insignis Herbivore Leaves/Stems not specific
Listronotus setosipennis Herbivore Stems to species Dhileepan and McFadyen, 2012; Strathie and McConnachie, 2013 Australia, South Africa
Lixus scrobicollis Herbivore Leaves not specific
Myrothecium roridum Pathogen not specific
Plasmopara halstedii Pathogen Leaves/Stems not specific
Platphalonidia mystica Herbivore Leaves not specific Dhileepan and McFadyen, 2012 Queensland
Podosphaera xanthii Pathogen Leaves not specific
Puccinia abrupta Pathogen not specific
Puccinia abrupta var. partheniicola Pathogen Inflorescence/Leaves to genus Dhileepan and McFadyen, 2012 Australia
Puccinia melampodii Pathogen Leaves not specific Dhileepan and McFadyen, 2012 Australia
Puccinia xanthii var. parthenii-hysterophorae Pathogen Leaves/Stems to genus Australia, South Africa
Smicronyx lutulentus Herbivore Seeds not specific Australia
Stobaera concinna Herbivore Leaves/Stems not specific
Trichoconiella padwickii Pathogen Leaves not specific Kaur and Aggarwal, 2015
Zygogramma bicolorata Herbivore Leaves not specific Dhileepan and McFadyen, 2012; Strathie and McConnachie, 2013 Australia; India; Karnataka; South Africa

Notes on Natural Enemies

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There is a considerable amount of literature concerning the natural enemies of P. hysterophorus, as the weed has been a target for biological control for more than 20 years. Between 1977 and 1991, extensive surveys of phytophagous arthropods were undertaken in its North American native range. Over 250 species were recorded during this period, and these have been assessed for specificity. The results of these studies have been summarized (McClay et al., 1995; Cock and Seier, 1999; Seier and Djeddour, 2000).

Six stenophagous insect species, including a leaf feeder, leaf miner, stem borer, stem galler and seed feeder, were released in Australia as biological control agents, and an additional two species from Argentina were released later (Navie et al., 1996; Evans, 1997a). The pathogens associated with P. hysterophorus in the neotropics have also been surveyed and evaluated as biocontrol agents (Evans, 1987b; 1997a, b). All the insect species released in Australia are coevolved natural enemies, as are the pathogens currently being assessed, however, numerous ad hoc surveys and studies of both the arthropods and pathogens associated with parthenium weed in India (Kumar, 1998) and Ethiopia (Taye, 2002) have also been undertaken. These all represent adaptive or opportunistic natural enemies, most probably polyphagous or from related genera of Compositae. The results of these studies have been fully documented (Singh, 1997; Evans, 1997a; Kumar, 1998). More recently, Kaur et al. (2014) provided lists of insects used as biocontrol agents, and of pathogenic fungi used as bioherbicidals. Among the most used insects for the biocontrol of P. hysterophorus are the leaf-feeding beetle Zygogramma bicolorata and the stem-galling moth Epiblema strenuana, A rust fungus, Puccinia abrupta var. partheniicola is also widely used as a bioherbicide (Gnanavel, 2013).

A wide range of crop insects and diseases has been reported from parthenium weed both in the neotropics and in its exotic range (McClay et al., 1995; Evans, 1997a; Singh, 1997). For example, it appears to be an important secondary host of a beetle pest (Pseudoheteronyx sp.) of sunflower in Australia, of plant parasitic nematodes in the USA (Navie et al., 1996), as well as of a major polyphagous lepidopteran pest (Diacrisia obliqua [Spilartica obliqua]) in India (Evans, 1997a). Similarly, it has been reported as a reservoir of Xanthomonas campestris pv. Phaseoli [X. axonopodis pv. phaseoli], Pseudomonas solancearum [Burkholderia solanacearum], Tomato yellow leaf curl virus, Potato X virus and Potato Y virus in both Cuba and India (Evans, 1997a). It is also known to host the major parasitic weeds, Orobanche spp. and Cuscuta spp. in Ethiopia.

Means of Movement and Dispersal

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Natural Dispersal

P. hysterophorus achenes are reported as being wind dispersed and capable of traveling long distances (Navie et al., 1996; Taye, 2002). The seeds are also dispersed in still or flood waters (Bajwa et al., 2016).

Vector Transmission (Biotic)

The seeds of P. hysterophorus can be dispersed by domestic and feral animals (Bajwa et al., 2016).

Accidental Introduction

The seeds P. hysterophorus have been reported as moved accidentally via irrigation water, farm machinery, industrial machinery, feral animals, humans, vehicles, stock fodder, movement of stock, grain and seed (PAG, 2000; Bajwa et al., 2016).

The transportation of soil, sand and gravel from P. hysterophorus-infested areas to non-infested areas for construction purposes may be the reason for the high infestation along roadsides and around buildings (Taye, 2002). Continental and inter-continental dispersal may occur when seeds contaminate commercial seed stocks or farm machinery. Seeds from the species have been found in large quantities in wash-down facilities used to clean vehicles and farm machinery. They have also been found in crop irrigation water and in mud attached to vehicles and farm animals (Bajwa et al., 2016).

Intentional Introduction

Human activities such as the use of P. hysterophorus as an ornamental plant, in floral bouquets, as packaging, and as a green manure, have contributed to its dispersal (Bajwa et al., 2016).

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Aid Yes
Animal production Yes Yes PAG, 2000
Crop production Yes Yes PAG, 2000
Cut flower trade Yes Yes
Disturbance Yes Yes
Escape from confinement or garden escapePossible from its use as an ornamental Yes
Flooding and other natural disasters Yes PAG, 2000
Forage Yes Yes PAG, 2000
Garden waste disposalPossible from its use as an ornamental Yes
HitchhikerAnimals, vehicles, machinery, grains, humans Yes Yes PAG, 2000; Baiwa et al., 2016
HorticultureOrnamental Yes Yes Baiwa et al., 2016
Medicinal useLocal ethnobotanical uses Yes Joshi et al., 2016
Military movementsAircraft and machinery parts during World War II. Yes Yes Auld et al., 1983
Seed trade Yes Yes PAG, 2000

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Bulk freight or cargo Yes PAG, 2000
Clothing, footwear and possessions Yes PAG, 2000
Containers and packaging - non-wood Yes Yes PAG, 2000
Land vehiclesTractors, combine harvestors, etc Yes
LivestockAttaches to animal fur Yes PAG, 2000
Machinery and equipment Yes Yes PAG, 2000
Plants or parts of plantsAnimal dung, compost, etc Yes Yes PAG, 2000
Soil, sand and gravelSoil, sand, irrigation water, flood Yes Shabbir et al., 2011; Taye, 2002
Water Yes PAG, 2000
Wind Yes Navie et al., 1996; Taye, 2002
Debris and waste associated with human activitiesAgricultural debris Yes Baiwa et al., 2016
Floating vegetation and debris Yes Baiwa et al., 2016
Pets and aquarium speciesAttached to domestic animals Yes Baiwa et al., 2016

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
Growing medium accompanying plants seeds Yes Pest or symptoms not visible to the naked eye but usually visible under light microscope
Stems (above ground)/Shoots/Trunks/Branches seeds Yes Pest or symptoms usually visible to the naked eye
True seeds (inc. grain) seeds Yes Pest or symptoms usually visible to the naked eye

Impact Summary

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CategoryImpact
Economic/livelihood Negative
Environment (generally) Negative
Human health Negative
Native fauna Negative

Economic Impact

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It is only in the past 20-30 years that P. hysterophorus has come to the fore as a weed of major economic importance, based mainly on its rapid spread in Australia and India. The impacts of parthenium weed were summarized by Parsons and Cuthbertson (1992), McFadyen (1992), Navie et al. (1996), Evans (1997a) and Mahadevappa (1997). Holm et al. (1991) recorded the species as a ‘serious’ weed in Mexico and a ‘principal’ weed in Cuba. Since its impact is multi-faceted, affecting crop production, animal husbandry, human health and biodiversity, its overall economic impact is difficult to quantify. The economic impact caused at various countries is due to the species outcompeting crops and pastures for resources, by the contamination of grains, by bringing pests and diseases into fields, and due to added eradication costs (Rubaba et al., 2017). The species is also responsible for causing the depreciation of land value in Africa, for which it is called ‘Faramsis’ in Ethiopia translating to ‘sign your land’ and ‘famine weed’ in South Africa.
 
The main impact of P. hysterophorus on crops relates to its allelopathic properties. The water soluble phenolics; caffeic acid, ferulic acid, vanicillic acid, anisic acid and fumaric acid; and sesquiterpene lactones, mainly parthenin and/or hymenin, occur in all parts of the plant and significantly inhibit the germination and subsequent growth of a wide variety of crops including pasture grasses, cereals, vegetables, other weeds and tree species (Navie et al., 1996; Evans, 1997a). Few critical assessments of yield losses have been made, although it has been determined that almost 30% grain loss can occur in irrigated sorghum in India (Channappagoudar et al., 1990). As Parthenium pollen is also allelopathic (Kanchan and Jayachandra, 1980), heavy deposits on nearby crop plants may result in failure of seed set, and losses of up to 40% have been reported in maize yield in India (Towers et al., 1977). In eastern Ethiopia, parthenium weed is the second most frequent weed (54%) after Digitaria abyssinica (63%) (Tamado and Milberg, 2000) and sorghum grain yield was reduced from 40 to 97% depending on the year and location (Tamado, 2001). Although P. hysterophorus is not yet considered to be a major crop weed in Australia (Navie et al., 1996), it has started to spread into sorghum, sugarcane and sunflower growing areas and negatively affect yields (Parsons and Cuthbertson, 1992). Also, Chippendale and Panetta (1994) estimate that cultivation costs may be doubled since the prepared ground has to be re-worked to eliminate the emergent parthenium weed seedlings.

Parthenium weed has also invaded forest land (Towers et al., 1977). It has become a menace in forest nurseries (Chandras, 1970). Swaminathan et al. (1990) reported its allelopathic effect on the multipurpose tree species Acacia leucocephala, Casuarina equisetifolia, Eucalyptus tereticornis and Leucaena leucocephala, along with some arable crops. The growth and nodulation of legumes were inhabited by parthenium weed because of the effect of allelochemicals on nitrogen fixing and nitrifying bacteria (Kanchan and Jayachandra, 1981; Dayama, 1986).

Another, indirect effect of P. hysterophorus on crop production is its role as an alternate host for crop pests. A wide range of crop insects and diseases has been reported from parthenium weed both in the neotropics and in its exotic range (McClay et al., 1995Evans, 1997a; Singh, 1997). For example, it appears to be an important secondary host of a beetle pest (Pseudoheteronyx sp.) of sunflower in Australia, of plant parasitic nematodes in the USA (Navie et al., 1996), as well as of a major polyphagous lepidopteran pest (Diacrisia obliqua [Spilartica obliqua]) in India (Evans, 1997a). Similarly, it has been reported as a reservoir of Xanthomonas campestris pv. Phaseoli [X. axonopodis pv. phaseoli], Pseudomonassolancearum [Burkholderia solanacearum], Tomato yellow leaf curl virus, Potato X virus and Potato Y virus in both Cuba and India (Evans, 1997a). It is also known to host the major parasitic weeds, Orobanche spp. and Cuscuta spp. in Ethiopia.

P. hysterophorus also significantly impacts on livestock production by affecting grazing land, animal health, milk and meat quality and the marketing of pasture seeds and feed grain. It can reduce the percentage cover of palatable species of grasslands in India by up to 90% (Jayachandra, 1971). The most comprehensive economic analysis has been made in Australia, where Parthenium weed monocultures in grazing land in Queensland were estimated to cover more than 17,000 km², reducing cattle stocking rates by as much as 80% (McFadyen, 1992), with a net annual loss of revenue calculated at up to AU$17 million (Chippendale and Panetta, 1994). Further losses result if farms also supply harvesting machinery, fodder or grain, since there is now legislation to prevent their movement from infested properties because of contamination by weed seed. An additional, non-quantifiable side effect of parthenium weed is on animal health, as the sesquiterpene lactone, parthenin, has been shown to cause severe dermatitis, anorexia and intestinal damage, which can lead to death of buffalo, cattle and sheep (Towers and Subba Rao, 1992), and 10-50% of the weed in the diet can kill these animals within 30 days (Naarasimhan et al., 1977a, b, 1980; More et al., 1982). Taints of meat have been detected from sheep given a diet of 30% parthenium weed (Tudor et al., 1982) and tainting of milk, meat and honey have also been reported (Towers and Subba Rao, 1992Taye, 2002).

Environmental Impact

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Impact on Habitats

Parthenium weed lacks predators, and cattle and livestock usually do not feed on it. As a result, the food chain is disturbed and the trophic structure changes, leading to an ecological imbalance in the invaded area. The importance value index (IVI) of parthenium weed remained at a maximum in both cropped and non-cropped areas across the seasons (Tiwari and Bisen, 1984). It causes a prolonged toxic effect to the soil environment – for instance, Kanchan and Jayachandra (1981) reported that the leachates from parthenium weed have an inhibitory effect on nitrogen fixing and nitrifying bacteria.

Parthenium weed is also an environmental weed that can cause irreversible habitat changes in native grasslands, woodlands, river banks and floodplains in both India and Australia (Jayachandra 1971McFadyen, 1992Evans, 1997aKumar and Rohatgi, 1999). In Nepal, the impacts include altering the soil nutrient composition and outcompeting the native plant species (India Biodiversity Portal, 2018). Huge stands of parthenium weed are common in almost all open areas. Parthenium weed, due to its allelopathic potential, replaces dominant flora and suppresses natural vegetation in a wide range of habitats and thus becomes a big threat to biodiversity. Batish et al. (2005) recorded 39 plant types in a Parthenium-free area, but only 14 were present in an infested area, and very little or sometimes no vegetation can be seen in some Parthenium-dominated areas (Kohli, 1992). Wherever it invades, it forms a territory of its own, replacing indigenous grasses and weeds, which are supposedly useful for the grazing animals (De and Mukhopadhyay, 1983). Parthenium weed has an adverse effect on a variety of natural herbs, which are the basis of traditional systems of medicines for the treatment of several diseases in various parts of the world (Mahadevappa et al., 2001Shabbir and Bajwa, 2006).

Impact on Biodiversity

P. hysterophorus has a detrimental effect on the health of grazing animals. Animals feeding on the species develop severe dermatitis, changes in their behavior, and lesions in the gastrointestinal tract, liver and kidney that can lead to death. Chemicals produced by the species that are washed away by water can get into aquatic ecosystems and have an adverse effect on aquatic plants. P. hysterophorus also affects the soil microbial activity (Joshi et al., 2016).

Ayele et al. (2013) did an assessment of the impact of P. hysterophorus on the above-ground vegetation and on the soil seed bank of herbaceous communities in Ethiopia. They found a 62.7% decrease of grass cover at the weed-infested sites. Also, the seeds of P. hysterophorus accounted for 84.2% of the overall seedbank in the highly infested areas. They concluded that the infestation of P. hysterophorus significantly reduced the amount and composition of both the above ground and the seed bank of herbaceous vegetation. Similar results have been reported for Australia by Nguyen et al. (2017): the species richness, diversity and evenness were all found to be significantly reduced as the density of P. hysterophorus increased. The authors also report the displacement of several native species.

Social Impact

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P. hysterophorus is an annual herb that is a poisonous or lethal weed for agricultural labourers and city-dwellers who are sensitive to it (Mukhopadhyay, 1987). Unfortunately, there is no effective treatment for the different kinds of allergies, other than avoiding contact or leaving the area. The impact of parthenium weed on human health in India (Lonkar et al., 1974Towers et al., 1977Towers and Mitchell, 1983; Towers and Subba Rao, 1992Kololgi et al., 1997), and more recently in Australia (McFadyen, 1995), is well documented. However, no economic analysis has been attempted despite reported widespread allergenic reactions to parthenium weed pollen and debris (trichomes). For example, 7% of the population in Bangalore (Karnataka State, India) suffers from allergenic rhinitis and over 40% is sensitive to the pollen (Towers and Subba Rao, 1992). Due to its chronic nature, there are reports of it leading to suicide in India (Kololgi et al., 1997). Also, Tanner and Mattocks (1987) hypothesised that Parthenium-contaminated animal feed leads to tainted milk, and that the hepatotoxic parthenin in tainted milk reacts synergistically with copper, causing Indian Childhood Cirrhosis (ICC). Deaths are reported by Rubaba et al. (2017) as a consequence of drinking tainted milk.

McFadyen (1995) predicted that after 1-10 years exposure to the weed, 10-20% of the population in newly infested areas in Australia would develop severe allergenic rhinitis, hence the cause for concern as parthenium weed spreads southwards into the more densely-populated urban areas of Queensland. A range of other, more serious health disorders have been attributed to parthenium weed, including allergenic eczematous contact dermatitis and bronchial asthma (Evans, 1997aKololgi et al., 1997). In Ethiopia, individuals hand weeding or hoeing in Parthenium-infested crops were reported to suffer from skin allergies, itching, fever and asthma (Taye, 2002). Affected individuals have no alternative except to leave the area. Therefore, it must be concluded that parthenium weed causes significant socio-economic damage in Parthenium-invaded countries. 

Risk and Impact Factors

Top of page Invasiveness
  • Invasive in its native range
  • Proved invasive outside its native range
  • Has a broad native range
  • Abundant in its native range
  • Highly adaptable to different environments
  • Is a habitat generalist
  • Pioneering in disturbed areas
  • Fast growing
  • Has high reproductive potential
  • Gregarious
  • Has propagules that can remain viable for more than one year
  • Has high genetic variability
Impact outcomes
  • Conflict
  • Damaged ecosystem services
  • Ecosystem change/ habitat alteration
  • Host damage
  • Modification of fire regime
  • Modification of successional patterns
  • Monoculture formation
  • Negatively impacts agriculture
  • Negatively impacts human health
  • Negatively impacts animal health
  • Negatively impacts livelihoods
  • Reduced native biodiversity
  • Damages animal/plant products
  • Negatively impacts trade/international relations
Impact mechanisms
  • Allelopathic
  • Causes allergic responses
  • Competition - monopolizing resources
  • Competition - shading
  • Pest and disease transmission
  • Induces hypersensitivity
  • Poisoning
  • Pollen swamping
  • Rapid growth
Likelihood of entry/control
  • Highly likely to be transported internationally accidentally
  • Difficult to identify/detect as a commodity contaminant
  • Difficult/costly to control

Uses

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Economic Value

Like many other weeds, parthenium weed also has beneficial aspects, and the search for a more effective utilization of parthenium weed will enable effective management on one hand and may provide productive uses instead of eradication on the other hand. Ramaswami (1997) and Seier and Djeddour (2000) reviewed the potential uses of parthenium weed. The application of parthenium weed compost and green leaf manure was reported to lower weed populations in rice. This was due to the role of allelopathic compounds present in it (Sudhakar, 1984), an increase in soil moisture due to the build-up of soil organic carbon (Son, 1995), increased soil N, P and K content (Bharati et al., 2001), and a reduced incidence of pests in rice such as stem borers and leaf rollers (Ramaswami, 1997). Parthenium weed is also potentially a rich source of potash (Parsons and Cuthbertson, 1992).
 

The allelopathic substances present in parthenium weed have been proposed as a source of insecticide (Parsons and Cuthbertson, 1992Hiremath and Ahn, 1997), herbicide (Mersie and Singh, 1987Pandy et al., 1993Batish et al., 2002), fungicide (Ganeshan and Jayachandra, 1993), and nematicide (Azam et al. 2001Prasad et al., 2002). Dwivedi et al. (2000) and Sharma et al. (2003) reported that parthenium weed extract has 95% repellency and oviposition deterrent properties against Callosobruchus chinensis in chick pea grains. A larval mortality of greater than 50% has been recorded in 2% concentration for Helicoverpa armigera (Sundararajan, 2002) and a significant decrease in life span and progeny production of the mustard aphid, Lipaphis erysimi, was reported (Sohal et al., 2002). Extract of parthenium weed was also found to significantly inhibit the growth of bacterial spot pathogen (Xanthomonas axonopodis pv. vesicatoria) infecting Capsicum frutescens (Sree and Sreeramulu, 2002).

Additional uses include as a foliar supplementation of the leaf water extract of parthenium weed on mulberry leaves stimulated silkworms to feed and utilize them more efficiently, resulting in vigorous growth of larvae, pupa cocoons and silk yield (Patil, 1997Singhal et al., 1998). Production of oxalic acid (Mane et al., 1986) and biogas (Gunaseelan, 1987Abubacker et al., 1999Thakur and Singh, 2000, 2003) from parthenium has been reported. In India, it is considered to be an exploitable source of easily extractable, high quality protein for stock feed (Savangikar and Joshi, 1978).

Social Benefit

P. hysterophorus is an annual herb used in its native neotropics as a herbal remedy for various intestinal and skin disorders using a decoction of boiled roots (Dominguez and Sierra, 1970). There are several reports on the antiviral, antifungal, antibacterial, antihelmintic and anti-inflammatory properties of the species (Joshi et al., 2016). It is used for the treatment of wounds, ulcerated sores, fever, anaemia, heart problems and malaria. It has potential medicinal properties due to antitumor (Mew et al., 1982) and antiamoebic activities (Sharma and Bhutani, 1988). It is also reported as used for toothaches, to remove boils and pimples, to relieve constipation, fevers, against insomnia and to treat diabetes (Joshi et al., 2016).

Environmental Services

The dry leaf powder of P. hysterophorus causes wilting of Salvinia molesta, which is an invasive species in some water bodies (Joshi et al., 2016). The species can be used for bioremediation of lead and nickel contaminated soils and water sources. The parthenin produced by the species has allelopathic effects, but also can stimulate growth of other plant species at subtoxic doses (Belz, 2016).

Uses List

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Environmental

  • Agroforestry
  • Soil improvement

Fuels

  • Biofuels

Materials

  • Chemicals
  • Green manure
  • Mulches
  • Pesticide
  • Poisonous to mammals

Medicinal, pharmaceutical

  • Source of medicine/pharmaceutical
  • Traditional/folklore

Ornamental

  • Cut flower
  • Potted plant

Similarities to Other Species/Conditions

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P. hysterophorus may be confused some Ambrosia species: A. artemisiifolia (annual ragweed), A. psilostachya (perennial ragweed), A. confertiflora (burr ragweed) and A. tenuifolia (lacy ragweed), especially during the rosette or vegetative stage. It can be distinguished from these species, which have opposite leaves in the early stages of growth, by the longitudinally-grooved stem. At the flowering stage, there can be no confusion, since the small white flower heads of P. hysterophorus, borne in much-branched terminal panicles, are readily distinguished from the greenish, spike-like racemes of Ambrosia (BioNet-EAFRINET, 2018).

Prevention and Control

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Prevention

As parthenium weed seed can be spread via flowing water or can be blown by the wind, the prevention of spread might be difficult. Seed production should be prevented by destroying the plants before flowering or seed setting. The seed can also stay for years in the soil seed bank and the continuous removal of the weed is required until the seed bank is depleted.

The spread of seed through the trading and transport of goods, animals grazing on infested fields, and the transportation of sand, soil and compost from infested areas to uninfested areas, are potential risks for further spread and hence should be controlled through quarantine.

In wastelands and public places such as gardens, recreation areas, parks, roadsides and railway tracks, where it is most serious, control has to be attempted on a community basis by local administrations. In India, campaigns have been attempted using children, the general public, NGOs and others (Bahn et al., 1997Singh, 1997), and in Ethiopia there was a wide publicity campaign through TV, radio, posters and seminars to bring awareness and educate people (Taye, 2002). Rubaba et al. (2017) suggest investing in surveillance systems in areas or countries of Africa where P. hysterophorus has not yet invaded to do early interventions. One important part of their proposed plan is training the general public in invasive species identification and providing invasive species fact sheets.

Parthenium weed is declared noxious throughout Queensland, Australia (Navie et al., 1996). It is categorized as P2 (where it must be destroyed) throughout the whole state except in specified areas where it is designated P3 and P4 (where infestations are to be reduced and prevented from spreading). In Queensland, parthenium weed seed is declared under the Agricultural Standards Act, which prevents the sale of commercial seed containing prohibited seed (Genn, 1987). Legislation has also been enacted to prevent the movement of vehicles carrying parthenium weed seed from Queensland to New South Wales, Australia and penalties have been imposed to deter the illegal entry of such vehicles (Parsons and Cuthbertson, 1992). It is a declared noxious weed in South Africa (Henderson, 2001). However, such prevention and quarantine activities are not exercised in many other countries and the weed is progressing in its invasion from continent to continent and country to country.

SPS Measures

Quarantine and eradication measurements are in effect for P. hysterophorus in Europe (EPPO, 2018). Australian authorities have imposed strict quarantines on contaminated equipment and stock to avoid spread into new areas (PIER, 2018).

Control

Eradication

In Kenya, the Suppression of Noxious Weeds Act of 2010 (CAP 325) obliges land owners to remove the species from their properties (Rubaba et al., 2017). The species is regulated as a quarantine pest in Europe under the EU IAS Regulation introduced in 2014 (EPPO, 2018), restricting its sale and movement, and requiring government agencies to undertake eradication programs.

Physical/mechanical control

Kaur et al. (2014) recommends the manual uprooting of the plants before flowering and seed set, followed by sowing desirable crops or pasture species. In some African countries with light infestations where labour is not too expensive, the use of machetes, hand pulling and burning of the species are a common practice (Rubaba et al., 2017). Physical removal by hand-pulling poses health risks and has not been recommended in Australia (Parsons and Cuthbertson, 1992). Mechanical treatments, such as grading, mowing, slashing and ploughing are also considered inappropriate since they may promote seed dispersal as well as rapid regeneration from lateral shoots close to the ground (Gupta and Sharma, 1977Navie et al., 1996). Fire has been used to control the first flush of emergent weeds at the beginning of the rains in Australia but is only considered to be a short-term control measure (Holman, 1981). A study by Vogler et al. (2002) showed that fire created open niches in the landscape, into which larger number of parthenium seeds were able to germinate in the absence of vegetation. Therefore, management of parthenium weed in pastures through burning is not considered to be an option.

Biological control

The use of insect and fungal pathogens and the exploitation of allelopathic plants is considered by Kaur et al. (2014) as the most economical and practical way to manage the infestations of the species. Biological control has been, and continues to be, considered the best long-term or sustainable solution to the parthenium weed problem in Australia (Haseler, 1976; McFadyen, 1992) and because of the vast areas and the socio-economics involved, this approach has also been proposed for India (Singh, 1997). South Africa was the first country in Africa to implement a biological control program against the species in 2003 (Rubaba et al., 2017). Four host-specific biocontrol agents have been released sequentially since 2010 after evaluation of their suitability, with variable establishment and spread (Strathie et al., 2016).

The use of insects as biocontrol agents had been tried in various countries (Kaur et al., 2014). Searches for, and evaluation of, coevolved natural enemies have been conducted in the neotropics since 1977. So far, nine insect species and two fungal pathogens have been introduced into Australia as classical biological control agents (Julien, 1992McClay et al., 1995; Navie et al., 1996; Dhileepan and McFadyen, 1997Evans, 1997a). Callander and Dhileepan (2016) report that most of these agents have become established and have proven effective in central Queensland, but that the weed is now spreading further into southern Queensland where the biocontrol agents are not present. Several of the agents are therefore now being redistributed into south and southeast Queensland.

The rust fungus, Puccinia abrupta var. partheniicola, is a prominent natural enemy in the semi-arid uplands of Mexico (Evans, 1987a, b), but since its release in Queensland in 1992, climatic conditions have been largely unfavourable (Evans, 1997a, b). It was accidentally introduced into Kenya (Evans, 1987a) and Ethiopia in mid-altitudes (1400-2500 masl) with disease incidence up to 100% in some locations (Taye et al., 2002a). Screening of another rust species (Puccinia melampodii) from Mexico was carried out (Evans, 1997b; Seier et al., 1997) and released in Australia in the summer of 1999/2000 (PAG, 2000). This fungus was later renamed Puccinia xanthii Schwein. var. parthenii-hysterophorae Seier, H.C.Evans & Á.Romero (Seier et al., 2009). Retief et al. (2013) report on specificity testing carried out in quarantine facilities in South Africa, and conclude that the fungus is suitable for release as a biological control agent of P. hysterophorus in South Africa. The authors suggest that this species has more potential for biocontrol in South Africa than Puccinia abrupta, which may have little impact in the low-altitude, high-temperature areas of the country where the weed is spreading.

In India, the mycoherbicide potential of plurivorous fungal pathogens belonging to the genera Fusarium, Colletotrichum, Curvularia,Myrothecium and Sclerotium, has and is being evaluated (Mishra et al., 1995; Evans, 1997a). Parthenium phyllody disease caused by the phytoplasma of faba bean phyllody group (FBP) was reported to reduce seed production by 85% (Taye et al., 2002b) and is being evaluated for use as a biological control agent in Ethiopia. Kaur and Aggarwal (2017) have tested an Alternaria isolate found on the weed, and report that it is worth investigating as a mycoherbicide for control of parthenium. Metabolites of Alternaria japonica and filtrates of Alternaria macrospora have caused significant damage to Parthenium (Kaur et al., 2015; Javaid et al., 2017).

Among the established insect biocontrol agents, the leaf-feeding beetle, Zygogramma bicolorata, the stem-galling moth, Epiblema strenuana, the stem-boring beetle, Listronotus setosipennis, and the seed-feeding weevil, Smicronyx lutulentus, are proving to be the most successful when climatic factors are favourable (McFadyen, 1992Dhileepan and McFadyen, 1997; Evans, 1997a). Some control of parthenium weed has also been achieved in India with Z. bicolorata (Jayanth and Visalakshy, 1994Singh, 1997Sarkate and Pawar, 2006), although there has been controversy concerning its taxonomy and host specificity (Jayanth et al., 1993Singh, 1997). Shabbir et al. (2016) reported that Z. bicolorata was most effective when applied in higher densities and at early growth stages of the weed. The distribution of this leaf beetle in South Asia was investigated by Dhileepan and Senaratne (2009), when it was present in many states in India, and in the Punjab region of Pakistan. Shrestha et al. (2011) reported that Z. bicolorata arrived in the Kathmandu Valley of Nepal in August 2010, and that by September it had spread over half of the valley areas where P. hysterophorus was present, although damage to the weed was only appreciable at one site.

Z. bicolorata has been seen attacking sunflowers in India and the use of Epiblema strenuata has not been effective, as it was found affecting Guizotia abyssinica crops (Kaur et al., 2014). More recently, Z. bicolorata and L. setosipennis have been released in South Africa and S. lutulentus is being evaluated under quarantine. Before approval as a biocontrol agent in South Africa in 2013, extensive testing suggested that Z. bicolorata would not become a pest of sunflowers in the country (McConnachie, 2015).

The use of antagonistic, competitor plants, such as Cassia spp. and Tagetes spp., has been recommended to control and replace P. hysterophorus in India (Mahadevappa and Ramaiah, 1988Evans, 1997aMahadevappa, 1997; Singh, 1997). In Australia, Bowen et al. (2007) tested a number of grass and legume species against the growth of parthenium weed plants and identified further species that could suppress weed growth. Recently, Khan et al. (2013) tested a number of native and introduced pasture species and identified several of them to be suppressive against parthenium weed in both glasshouse and field conditions. The sowing of selected pasture plants in infested areas can suppress the growth of parthenium weed by as much as 80% and also provide improved fodder for stock (Adkins et al., 2012). Shabbir et al. (2013) showed that the suppressive plants and biological control agents can act synergistically to significantly reduce both the biomass and seed production of parthenium weed under field conditions. The suppressive plants strategy is easy to apply, sustainable over time, profitable under a wide range of environmental conditions and promotes native plant biodiversity. Species reported as effectively outcompeting P. hysterophorus are Cassia sericea, C. tora, C. auriculata, Croton bonplandianum, Amaranthus spinosus, Tephrosia purpurea, Hyptis suaveolens, Sida spinosa, and Mirabilis jalapa. Extracts of Imperata cylindrica, Desmostachya bipinnata, Otcantium annulatum, Sorghum halepenseDicanthium annulatum, Cenchrus pennisetiformis, Azadirachta indica, Aegle marmelos and Eucalyptus tereticornis are reported as inhibiting the germination and/or growth of P. hysterophorus (Kaur et al., 2014).

Chemical control

The chemical control of Parthenium hysterophorus is effective in areas lacking natural enemies, specially at the rosette stage (Kaur et al., 2014). A range of herbicides including atrazine, dicamba, 2,4-D, picloram and glyphosate, all applied at high volume, have been employed successfully in Queensland, Australia (Haseler, 1976). However, chemical control over the enormous areas infested by parthenium weed in Queensland is economically unviable and non-sustainable (Parsons and Cuthbertson, 1992), as well as environmentally undesirable (Navie et al., 1996). In India, the economics of spraying are even more untenable. Nevertheless, in Australia, spot spraying with atrazine plus a non-ionic surfactant is recommended as a pre-emergence treatment. Post-emergence control has been achieved with 2,4-D, often in combination with picloram (Navie et al., 1996), whilst low rates of glyphosate have proven to be effective in coffee plantations in Kenya (Njoroge, 1989).

Some of the newer herbicides, such as imazapyr, oxadiazon, oxyfluorfen, pendimethalin and thiobencarb, have also been reported to be highly effective against parthenium weed (Parsons and Cuthbertson, 1992). Imazethapyr is particularly effective as a pre-emergence treatment in green gram (Tewari et al., 2004). Bromoxynil + MCPA was the most effective of a range of post-emergence treatments tested by Javaid (2007). Glyphosate, glufosinate, chlorimuron and trifloxysulfuron applied at the rosette stage provided greater than 93% control, while halosulfuron, MSMA, bromoxynil, 2,4-D, and flumioxazin gave 58-90% control (Reddy et al., 2007), and norflurazon and clomazone were also highly effective.

There are several disadvantages for the use of chemical herbicides, such as the development of resistance to some herbicides and having detrimental effects over other species. For example, glyphosate is very toxic and can be damaging to the wild flora (Kaur et al., 2014). Lorenzi (1984) indicates susceptibility to acifluorfen, ametryne, atrazine, bentazon, bifenox, cyanazine, dicamba, diquat, diuron, 2,4-D, fomesafen, glyphosate, ioxynil, linuron, metribuzin, molinate, napropamide, oxadiazon, oxyfluorfen, paraquat, prometryne, simazine and tebuthiuron, while there is moderate to total resistance to alachlor, asulam, butachlor, butylate, EPTC, oryzalin, pendimethalin, trifluralin and vernolate. However, in Brazil, herbicide resistance has developed in relation to the ALS-inhibiting herbicides, i.e. imidazolinones (imazethapyr), triazolopyrimidines (cloransulam-methyl), sulfonylureas (chlorimuron-ethyl and iodosulfuron-methyl-sodium plus foramsulfuron. In these areas, 2,4-D is used as an alternative (Gazziero et al., 2006). Atrazine is restricted as some countries due to its groundwater contamination potential (Kaur et al., 2014).

IPM

In many locations parthenium weed is able to survive individually-applied management measures, and a more effective integrated approach is therefore required in these locations. A holistic IPM approach is propounded in India to achieve sustainable management of parthenium weed (Mahadevappa, 1997), and implemented in Australia through improved extension strategies (Navie et al., 1996Chamala et al., 1997). Nav-Bahr and Bahar (2000) proposed ploughing before flower set and burning when the plants are dry and mature, application of atrazine or other herbicides like 2,4-D, paraquat, glyphosate diuron and dalapon, using Cassia sericea to displace parthenium weed, and biocontrol using Zygograma bicolorata.

Gaps in Knowledge/Research Needs

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Several research needs have been identified by various authors regarding P. hysterophorus. There is limited knowledge of the species impact on the agriculture, biodiversity, and human and animal health in Africa. No mapping for the presence of the species is available for several countries such as Zambia, Swaziland, Botswana and Zimbabwe. Therefore, the actual extent of the spread of P. hysterophorus in Africa is not known. Studies on how climate change might influence the growth and survival of the species are needed. Studies on the contribution of genetic diversity in the species invasion mechanism are also needed (Bajwa et al., 2016).

Research and development priorities concerning the species management, health impacts, and its possible benefits are also recommended by Rubaba et al. (2017).

References

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23/02/2018 Updated by:

Jeanine Vélez-Gavilán, University of Puerto Rico at Mayagüez

30/09/13 Updated by:

Asad Shabbir, University of the Punjab, Pakistan

15/04/08 Updated by:

Chris Parker, Consultant, UK

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