Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Rottboellia cochinchinensis
(itch grass)

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Datasheet

Rottboellia cochinchinensis (itch grass)

Summary

  • Last modified
  • 08 November 2018
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Host Plant
  • Preferred Scientific Name
  • Rottboellia cochinchinensis
  • Preferred Common Name
  • itch grass
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Plantae
  •     Phylum: Spermatophyta
  •       Subphylum: Angiospermae
  •         Class: Monocotyledonae
  • Summary of Invasiveness
  • The erect, profusely tillering annual grass R. cochinchinensis grows up to a height of 4 m or more and is extremely competitive with annual crops, readily invading disturbed sites along roads and railways. Co...

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Pictures

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PictureTitleCaptionCopyright
Mature R. cochinchinensis weed (at least 5 m high) in sorghum, Nigeria.
TitleMature weed
CaptionMature R. cochinchinensis weed (at least 5 m high) in sorghum, Nigeria.
Copyright©Chris Parker/Bristol, UK
Mature R. cochinchinensis weed (at least 5 m high) in sorghum, Nigeria.
Mature weedMature R. cochinchinensis weed (at least 5 m high) in sorghum, Nigeria.©Chris Parker/Bristol, UK
a, Ligule, ventral view; b, part of raceme; c, joint of raceme with grain; d, lower glume (G1), dorsal view (opened); e, upper glume (G2), lateral view; f, lower lemma (L1), dorsal view; g, lower palea (P1), ventral view; h, upper lemma (L2), lateral view; i, upper palea (P2), dorsal view; j, caryopsis.
TitleR. exaltata [R. cochinchinensis] - line drawing
Captiona, Ligule, ventral view; b, part of raceme; c, joint of raceme with grain; d, lower glume (G1), dorsal view (opened); e, upper glume (G2), lateral view; f, lower lemma (L1), dorsal view; g, lower palea (P1), ventral view; h, upper lemma (L2), lateral view; i, upper palea (P2), dorsal view; j, caryopsis.
CopyrightSEAMEO-BIOTROP
a, Ligule, ventral view; b, part of raceme; c, joint of raceme with grain; d, lower glume (G1), dorsal view (opened); e, upper glume (G2), lateral view; f, lower lemma (L1), dorsal view; g, lower palea (P1), ventral view; h, upper lemma (L2), lateral view; i, upper palea (P2), dorsal view; j, caryopsis.
R. exaltata [R. cochinchinensis] - line drawinga, Ligule, ventral view; b, part of raceme; c, joint of raceme with grain; d, lower glume (G1), dorsal view (opened); e, upper glume (G2), lateral view; f, lower lemma (L1), dorsal view; g, lower palea (P1), ventral view; h, upper lemma (L2), lateral view; i, upper palea (P2), dorsal view; j, caryopsis.SEAMEO-BIOTROP
The inflorescence is a cylindrical raceme that is 3-15 cm long. Picture taken from Ethiopian specimen.
TitleInflorescence (detail)
CaptionThe inflorescence is a cylindrical raceme that is 3-15 cm long. Picture taken from Ethiopian specimen.
Copyright©Chris Parker/Bristol, UK
The inflorescence is a cylindrical raceme that is 3-15 cm long. Picture taken from Ethiopian specimen.
Inflorescence (detail)The inflorescence is a cylindrical raceme that is 3-15 cm long. Picture taken from Ethiopian specimen.©Chris Parker/Bristol, UK
Inflorescence infected by smut, Ethiopia.
TitleInflorescence infected by smut
CaptionInflorescence infected by smut, Ethiopia.
Copyright©Chris Parker/Bristol, UK
Inflorescence infected by smut, Ethiopia.
Inflorescence infected by smutInflorescence infected by smut, Ethiopia.©Chris Parker/Bristol, UK
R. cochinchinensis seeds on soil, Ethiopia.
TitleSeeds
CaptionR. cochinchinensis seeds on soil, Ethiopia.
Copyright©Chris Parker/Bristol, UK
R. cochinchinensis seeds on soil, Ethiopia.
SeedsR. cochinchinensis seeds on soil, Ethiopia.©Chris Parker/Bristol, UK

Identity

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Preferred Scientific Name

  • Rottboellia cochinchinensis (Lour.) Clayton

Preferred Common Name

  • itch grass

Other Scientific Names

  • Aegilops exaltata L.
  • Aegilops fluviatilis Blanco
  • Manisuris exaltata Kuntze
  • Manisuris exaltata var. appendiculata (Steud.) Honda
  • Ophiuros appendiculatus Steud.
  • Rottboellia arundinacea Hochst. ex A. Rich
  • Rottboellia denudata Steud.
  • Rottboellia exalata fo. arundinacea Hochst. ex A. Rich Hack.
  • Rottboellia exaltata L. f.
  • Rottboellia exaltata var. appendiculata (Steud.) Hack.
  • Rottboellia setosa J.S. Presl ex C.B. Presl
  • Stegosia conchinchinensis Lour.,
  • Stegosia exaltata Nash

International Common Names

  • English: corn grass; guineafowl grass; guineafowlgrass; itchgrass; jointed grass; kokoma grass; prickle grass; raoul grass; rice grass; shamvagrass; sugarcane weed; treadmill
  • Spanish: caminadora; cebada fina; graminea corredora
  • French: herbe a canne; herbe a riz; herbe queue-de-rat
  • Chinese: tong zhou mao
  • Portuguese: capim-camalote

Local Common Names

  • Brazil: grama-alta; rabo-de -lagarto
  • Costa Rica: zacate de fuego; zacate indio
  • Cuba: grama de caballo; sancarana
  • India: barsali; bura; dholu; konda panookoo; swooate
  • Indonesia: bandjangan; bayung; bludru; branjangan; doekoet kikisian; jukut kikisan
  • Japan: tsunoaiashi
  • Malawi: kadawe; kandulu
  • Philippines: agingai; anguigay; annarai; bodo; bukal; gaho; girum nagei; nagel; sagisi
  • South Africa: tarentaalgras
  • Thailand: yaa prong khaai
  • Venezuela: paja peluda
  • Zambia: mulungwe; shamwe grass
  • Zimbabwe: kokomo grass; shamva

EPPO code

  • ROOEX (Rottboellia exaltata)

Summary of Invasiveness

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The erect, profusely tillering annual grass R. cochinchinensis grows up to a height of 4 m or more and is extremely competitive with annual crops, readily invading disturbed sites along roads and railways. Commonly known as itchgrass, it has brace roots near the base of the plant, a cylindrical spikelet seedhead and siliceous hairs on the leaf sheath that can penetrate and irritate the skin. Individual plants produce 2000 to 16,000 seeds that are shed as soon as they mature. A native of Indo-China, it is naturalised throughout the tropics of Asia, and is found in north-eastern Australia and savannah zones of Africa. It has been introduced into tropical America, as a potential pasture grass in the USA in the early 1900s and since the 1960s has been spread widely by contaminated rice seed, agricultural equipment and along transport routes in Central and South America, the Caribbean, and in the Gulf Coast region of the USA. An aggressive, significant weed in more than 40 countries, R. cochinchinensis is listed as a Federal Noxious Weed in the USA, and is suggested by Vibrans (2009) to be possibly the most harmful invasive plant in Mexico.

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Plantae
  •         Phylum: Spermatophyta
  •             Subphylum: Angiospermae
  •                 Class: Monocotyledonae
  •                     Order: Cyperales
  •                         Family: Poaceae
  •                             Genus: Rottboellia
  •                                 Species: Rottboellia cochinchinensis

Notes on Taxonomy and Nomenclature

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Rottboellia is a small genus of species native to the tropics of Asia and widespread in Africa. It is a typical member of the grass tribe Andropogoneae, characterized by the inflorescence disarticulating into floral units consisting of a sessile spikelets, pedicellate spikelet, and internode. Some closely related genera include Manisuris, Coelorachis, and Hemarthria.

Rottboellia cochinchinensis (Lour.) W.D. Clayton is the accepted name of a species known until 1981 as R. exaltata L.f., an illegitimate name since it was already in use for a different species. . A subsequent proposal to retain the name R. exaltata was made (Simon, 1982), but was not  accepted, therefore R. conchinchinensis is the correct name to use for this species.
 

Description

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R. cochinchinensis is an erect annual grass that grows up to a height of 4 m or more. The inflorescence is a cylindrical raceme that is 3-15 cm long. The floral units consist of a sessile spikelet, pedicellate spikelet and internode. The pedicel is fused to the swollen floral internode. The spikelets are awnless, 3.5-6 mm long, and 2.5-3 mm wide. The floral units separate and fall as soon as they mature, from the top of the raceme downwards.

Stout, strongly tufted, frequently with aerial prop roots; flowering culms 0.5-3.3 m tall, branched, 5-19 noded, nodes glabrous; internodes glabrous, grooved, ribbed, solid to spongy. Leaves 15-60 cm long, 0.5-2.5 cm broad. Leaf sheaths keeled to rounded, ribbed, smooth, covered with long, sharp, silicaceous, tubercle-based, fragile, irritating hairs that break off on contact, upper sheaths glabrous or hairy, auricles absent. Ligule short, fringed with hairs, 1-2 mm long, truncate; blades flat, keeled, 20-60 cm long, 1-2.5 cm wide, linear-lanceolate, acuminate, base cordate, hairy or glabrous, scabrous, margins very rough.

Inflorescence, a jointed raceme at the terminus of the culm and each branch of the culm; spikes cylindrical, 8-15 cm long, about 3 mm in diameter, glabrous, sheathed at the base, readily breaking into hard cylindrical joints that are 6-7 mm long; spikelets sometimes sterile; sessile spikelets 5-7 mm long, as long as the joint or distinctly shorter; pedicelled spikelets 3-6 mm long; 5-8 mm long, deeply grooved on the lower part, apex hollow; callus soft, smooth, truncate and peg-like. Pedicel similar in appearance and fused to the internode, 3-5-6 mm long, 2.5-3 mm wide. First glume as long as the spikelet, oblong to lanceolate, 9-11 nerved, rounded to cleft, indurate, convex, muricate, glabrous, slightly winged at the apex, margins enrolled. Second glume many nerved, keeled on the upper part, boat-shaped, smooth on the lower part and muricate upwards, indurate to cartilaginous, following the outline of the internode. Lower lemma oblong to lanceolate, faintly 3-nerved, as long as the second glume, membraneous to chartaceous. Lower palea well developed and similar to the lemma. Upper floret perfect, hyaline. Caryopsis oblong, 3-4 mm long, 1.75-2.0 mm broad, face inflated and loosely wrinkled, back more or less flattened, golden brown with a conspicuous dark brown spot above the hilum, deeply embedded in the indurate rachis (Holm et al., 1977; Ivens et al., 1978; Webster, 2003).

Plant Type

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Grass / sedge
Seed propagated

Distribution

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R. cochinchinensis is a native of the Old World tropics, and is thought to have an origin in Indo-China in the area that is now Vietnam. It is now present in tropical areas of the Americas and Caribbean, as well as being widespread in tropical Asia and the Pacific Islands, tropical Africa, and Australia.

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

BangladeshPresentNativeClayton et al., 2014
CambodiaPresentNativeClayton et al., 2014
ChinaRestricted distributionUSDA-ARS, 2008; EPPO, 2014
-FujianPresentNativeFlora of China Editorial Committee, 2008
-GuangdongPresentNativeFlora of China Editorial Committee, 2008
-GuangxiPresentNativeFlora of China Editorial Committee, 2008
-GuizhouPresentNativeFlora of China Editorial Committee, 2008
-HainanPresentNativeFlora of China Editorial Committee, 2008
-Hong KongRestricted distributionEPPO, 2014
-HunanPresentNativeFlora of China Editorial Committee, 2008
-SichuanPresentNativeFlora of China Editorial Committee, 2008
-YunnanPresentNativeFlora of China Editorial Committee, 2008
-ZhejiangPresentNativeFlora of China Editorial Committee, 2008
Christmas Island (Indian Ocean)PresentNativeOrchard, 1993
IndiaRestricted distributionNativeChristopher et al., 1989; USDA-ARS, 2008; EPPO, 2014
-Andaman and Nicobar IslandsPresentNativeShukla, 1996
-Andhra PradeshPresentNativeShukla, 1996
-AssamPresentNativeShukla, 1996
-BiharPresentNativeShukla, 1996
-Himachal PradeshPresentNativeShukla, 1996
-Madhya PradeshPresentNativeShukla, 1996
-MeghalayaPresentNativeShukla, 1996
-NagalandPresentNativeShukla, 1996
-OdishaPresentNativeShukla, 1996
-Tamil NaduPresentNativeShukla, 1996
-Uttar PradeshPresentNativeShukla, 1996
-West BengalPresentNativeShukla, 1996
IndonesiaRestricted distributionNativeHolm et al., 1977; Waterhouse, 1993; USDA-ARS, 2008; EPPO, 2014
-JavaPresentGBIF, 2008
-Nusa TenggaraPresentGBIF, 2008
-SumatraPresentRNG, 2008
JapanPresentNativeNakama et al., 1988; USDA-ARS, 2014
-Ryukyu ArchipelagoPresentNativeUSDA-NRCS, 2008Ryukyu Islands
Korea, Republic ofPresentKim et al., 1993
KyrgyzstanPresentWaterhouse, 1993
LaosPresentNativeWaterhouse, 1993; USDA-ARS, 2014
MalaysiaRestricted distributionNativeReed et al., 1977; Waterhouse, 1993; USDA-ARS, 2008; EPPO, 2014
MyanmarRestricted distributionNativeWaterhouse, 1993; USDA-ARS, 2008; EPPO, 2014
NepalPresentNativeUSDA-ARS, 2008
OmanPresentNativeClayton et al., 2014
PakistanPresentCope, 1982Found in Punjab
PhilippinesRestricted distributionNativePamplona et al., 1976; Waterhouse, 1993; EPPO, 2014
SingaporePresentDuistermaat, 2004
Sri LankaPresentNativeHolm et al., 1977; USDA-ARS, 2008Specimens from North and North Central Provinces (GBIF, 2008)
TaiwanPresentNativeMillhollon and Burner, 1993; USDA-ARS, 2008
ThailandRestricted distributionNativeMillhollon and Burner, 1993; Waterhouse, 1993; USDA-ARS, 2008; EPPO, 2014
VietnamPresentNativeWaterhouse, 1993; Oviedo Prieto et al., 2012
YemenPresentNativeClayton et al., 2014

Africa

AngolaPresentNativeHolm et al., 1977; Clayton et al., 2014
BeninNativeLutzeyer and Koch, 1992; USDA-ARS, 2008
BotswanaNativeCope, 2002; USDA-ARS, 2008
Burkina FasoNativeUSDA-ARS, 2008
BurundiPresentNativeGBIF, 2008; Clayton et al., 2014
CameroonPresentNativeMartin, 1990; USDA-ARS, 2008
Cape VerdePresentNativeLima and Duclos, 2001; Clayton et al., 2014
Central African RepublicPresentNativeGBIF, 2008; Clayton et al., 2014Central Province
ChadPresentNativeClayton et al., 2014
Côte d'IvoireRestricted distributionUSDA-ARS, 2008; EPPO, 2014
Equatorial GuineaPresentNativeUSDA-ARS, 2008
EritreaPresentNativeClayton et al., 2014
EthiopiaRestricted distributionNativeHolm et al., 1977; USDA-ARS, 2008; EPPO, 2014
GambiaPresentNativeUSDA-ARS, 2008
GhanaRestricted distributionNativeHaizel, 1973; USDA-ARS, 2008; EPPO, 2014An important invasive weed in Ashanti region (Anning and Yeboah-Gyan, 2007)
GuineaRestricted distributionUSDA-ARS, 2008; EPPO, 2014
Guinea-BissauPresentNativeRNG, 2008; Clayton et al., 2014
KenyaRestricted distributionNativeMichieka, 1991; USDA-ARS, 2008; EPPO, 2014
LiberiaPresentNativeClayton et al., 2014
MadagascarRestricted distributionHolm et al., 1977; Clayton et al., 2014; EPPO, 2014
MalawiPresentBanda and Morris, 1986; Clayton et al., 2014Especially common in Dowa, Kasungu, Lilongwe and Mua Districts
MaliPresentNativeGBIF, 2008; Clayton et al., 2014
MozambiqueRestricted distributionHolm et al., 1977; Clayton et al., 2014; EPPO, 2014
NamibiaPresentNativeUSDA-ARS, 2008
NigeriaRestricted distributionNativeOkafar and Zitta, 1991; USDA-ARS, 2008; EPPO, 2014
RéunionPresentMillhollon and Burner, 1993
SenegalRestricted distributionClayton et al., 2014; EPPO, 2014
SeychellesPresentNativeClayton et al., 2014
Sierra LeonePresentAlghali and Domingo, 1982; USDA-ARS, 2008
SomaliaPresentNativeUSDA-ARS, 2008
South AfricaRestricted distributionNativeSouth African Sugar Association, 1980; Bromilow, 2001; USDA-ARS, 2008; EPPO, 2014
SudanRestricted distributionNativeHolm et al., 1977; USDA-ARS, 2008; EPPO, 2014
SwazilandNativeSwaziland National Trust Commission, 2008; USDA-ARS, 2008
TanzaniaRestricted distributionNativeMillhollon and Burner, 1993; EPPO, 2014Mbeya, Morogoro, Rukwa and Kigoma regions
-ZanzibarPresentRNG, 2008
TogoPresentNativeGBIF, 2008; Clayton et al., 2014
UgandaRestricted distributionNativeMillhollon and Burner, 1993; USDA-ARS, 2008; EPPO, 2014
ZambiaPresentNativeHolm et al., 1977; Clayton et al., 2014
ZimbabweRestricted distributionRambakudibga, 1988; EPPO, 2014

North America

MexicoPresentIntroducedGBIF, 2008; Vibrans, 2009Known in Oaxaca in 1955, specimens in 1970s and 1980, also from Campeche, Tabasco and Veracruz (GBIF, 2008)
USAWidespreadIntroduced Invasive Holm et al., 1977; EPPO, 2014
-AlabamaPresentIntroducedLorenzi and Jeffery, 1987; USDA-NRCS, 2014
-ArkansasPresent, few occurrencesIntroduced1982GBIF, 2008; USDA-ARS, 2008; USDA-ARS, 2014; USDA-NRCS, 2014In Arkansas and Ashley counties (Arkansas State Plant Board, 2008)
-CaliforniaPresentIntroducedUSDA-ARS, 2014
-FloridaPresentLorenzi and Jeffery, 1987; USDA-ARS, 2008; USDA-NRCS, 2014
-GeorgiaIntroduced Invasive USDA-ARS, 2008; USDA-ARS, 2014; USDA-NRCS, 2014
-IllinoisPresentIntroducedUSDA-ARS, 2014
-IndianaIntroduced Invasive USDA-ARS, 2008
-LouisianaPresentIntroduced1927Lorenzi and Jeffery, 1987; USDA-ARS, 2008In 1927 near Ruth, St. Martin Parish (Millhollon, 1975)
-MississippiPresentLorenzi and Jeffery, 1987; USDA-ARS, 2008; USDA-NRCS, 2014
-MissouriIntroducedGBIF, 2008In Missouri University Research Park in 1975 (GBIF, 2008)
-North CarolinaPresentMillhollon and Burner, 1993; USDA-ARS, 2008; USDA-NRCS, 2014
-South CarolinaIntroduced Invasive Webster, 2003
-TexasIntroduced1992 Invasive USDA-ARS, 2008; USDA-NRCS, 20141992, Jefferson County
-UtahPresentIntroducedUSDA-ARS, 2014

Central America and Caribbean

BahamasPresentIntroducedMissouri Botanical Garden, 2008
BarbadosPresentIntroducedBroome et al., 2007
BelizePresentIntroducedGBIF, 2008Cayo, Corozal and Orange Walk Provinces
Cayman IslandsPresentIntroducedAcevedo-Rodriguez and Strong, 2012
Costa RicaPresentIntroducedCruz et al., 1994; Chacón and Saborío, 2012; EPPO, 2014Specimen from Turrialba in 1949 but present in Cartago, Guanacaste and Limon Provinces from at least 1966 and Puntarenas from 1976. Also now in Alajuela, Heredia and San Jose (GBIF 2008)
CubaPresentIntroduced Invasive Gutierrez and Gonzalez, 1993; EPPO, 2014
Dominican RepublicPresentIntroducedUlloa et al., 1990; EPPO, 2014Potentially invasive (Kairo et al., 2003)
El SalvadorPresentIntroducedMissouri Botanical Garden, 2008
GrenadaPresentIntroducedBroome et al., 2007
GuadeloupePresentIntroducedMillhollon and Burner, 1993; Broome et al., 2007Specimen from 1963 (GBIF, 2008)
GuatemalaPresentIntroducedJiminez et al., 1990; EPPO, 2014
HaitiPresentIntroducedMissouri Botanical Garden, 2008
HondurasPresentIntroducedMillhollon and Burner, 1993; EPPO, 2014In Atlantida and Olacho state/province (GBIF, 2008)
JamaicaRestricted distributionIntroducedHolm et al., 1977; EPPO, 2014
MartiniquePresentIntroducedBroome et al., 2007
NicaraguaWidespreadIntroducedGBIF, 2008Pacific Coastal Plain in states of Chontales, Leon and Manuagua
PanamaRestricted distributionIntroducedMillhollon and Burner, 1993; EPPO, 2014Collected in 1960 from Canal Area. Subsequently from Bocas de Toro and Chiriqui states
Puerto RicoPresentVelez and Semidey, 1991; USDA-ARS, 2008; Rojas-Sandoval and Acevedo-Rodríguez, 2014
Saint Kitts and NevisPresentIntroducedMissouri Botanical Garden, 2008Nevis
Saint LuciaPresentIntroducedBroome et al., 2007
Saint Vincent and the GrenadinesPresentIntroducedBroome et al., 2007
Trinidad and TobagoRestricted distributionIntroducedHolm et al., 1977; EPPO, 2014

South America

ArgentinaPresentIntroducedMillhollon and Burner, 1993
BoliviaRestricted distributionIntroducedUnterladstatter, 1979; EPPO, 2014
BrazilRestricted distributionIntroducedMillhollon and Burner, 1993; EPPO, 2014
-Espirito SantoPresentIntroducedGBIF, 2008
-Mato GrossoPresentIntroducedFilgueiras and Valls, 2014Naturalized
-Mato Grosso do SulPresentIntroducedFilgueiras and Valls, 2014Naturalized
-Rio de JaneiroPresentIntroducedFilgueiras and Valls, 2014Naturalized
-RoraimaPresentIntroducedFilgueiras and Valls, 2014Naturalized
-Sao PauloPresentIntroducedLorenzi, 1982
ColombiaRestricted distributionIntroducedHolm et al., 1977; EPPO, 2014Collected in Dept. El Valle, Palmira in 1963. Known from Antioquia and Valle de Cauca Provinces (GBIF, 2008)
EcuadorPresentIntroducedCarcelen, 1975; Millhollon and Burner, 1993Collected from Chimborazo, Cotopaxi, Galapagos, Guayas and Pichincha state/province (GBIF, 2008)
-Galapagos IslandsPresentIntroduced Invasive Charles Darwin Foundation, 2008Invasive on Santa Cruz Island
French GuianaPresentIntroducedClayton et al., 2014
PeruRestricted distributionIntroduced Invasive Pleasant et al., 1990; EPPO, 2014Huanuco, Loreto, San Martin and Ucayali states/provinces
SurinameIntroducedGBIF, 2008Collected in 1956
VenezuelaRestricted distributionIntroducedHolm et al., 1977; EPPO, 2014Specimen from 1966 Acariqua (GBIF, 2008). Also from Amazonas, Cojeded and Zulia provinces

Oceania

AustraliaRestricted distributionHolm et al., 1977; EPPO, 2014
-Australian Northern TerritoryPresentNativeUSDA-ARS, 2008
-New South WalesPresentNativeUSDA-ARS, 2008
-QueenslandPresentNativeUSDA-ARS, 2008
FijiPresentNativeWaterhouse, 1997
Papua New GuineaPresentNativeVance, 1982Morobe and Western Provinces (GBIF, 2008)
Solomon IslandsPresentNativeSwarbrick, 1997

History of Introduction and Spread

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R. cochinchinensis was probably introduced to the New World at the beginning of the 1900s, and now occurs in tropical areas of Central and Southern America, the Caribbean, and in the Gulf Coast region of the USA (Holm et al., 1977; Strahan et al., 2000). It is widespread in tropical Asia and the Pacific Islands, tropical Africa, and Australia. It is common in many parts of West Africa, especially in fallows and waste land in the savannah zone (Ivens et al., 1978). In Central America and the Caribbean it is estimated that itchgrass affects more than 3.5 million hectares (FAO, 1992). In the Caribbean region, this species first appears in collections made in 1909 in Cuba and 1912 in Jamaica (US National Herbarium). 

R. cochinchinensis was first introduced into the USA from the Philippines in the early 1900s as a potential pasture grass (Bugwood, 2008), and into the southern USA early in the 1900s via the West Indies, in plant material or on equipment (Millhollon, 1965), by birds (Aison et al., 1984), flood water, rodents, and farm machinery (Millhollon, 1980; Freshwater et al. 1986; Strahan et al., 2000a). The spread of R. cochinchinensis in Louisiana has been attributed in part to movement of equipment and road matting materials used during the extensive oil exploration that occurred in the late 1970s and early 1980s, and by 1980, the weed infested 38 Louisiana parishes on about 80,940 ha (Millhollon, 1980). Machinery was also responsible for introduction into Arkansas, first discovered in September 1982 on a farm in Ashley County from seed carried in crop debris on a combine harvester used in previous harvests in Louisiana (Arkansas State Plant Board, 2008). It has also been reported that movement into new areas has been along railroad tracks, and its appearance in south-eastern North Carolina in the early 1980s along the north-south seaboard coastline railroad is an example of this mode of spread (Hall and Patterson, 1992). The species is one of the most frequently intercepted Federal noxious weeds found by inspectors at the USA border, as seeds often hitchhike on railroad cars from Mexico (Lehtonen, 2003).

Introductions have also been through contaminated crop seed for trials, exchanged by international research programmes or in commercial seed supplied to farmers. For example Rottboellia has been found in rice seed lots received at the International Rice Research Institute in the Philippines (Huelma et al., 1996). Similarly, there are indications of dissemination in rice seed movements from Colombia to Brazil in 1961 (Millhollon and Burner, 1993). In Campeche and other areas in Mexico, farmers identified contaminated rice seed as being responsible for the introduction of R. cochinchinensis to their fields (Valverde et al., 1999b, 2001). Movement of livestock and farm machinery is blamed for the introduction of an exotic biotype from Thailand to Malaysia (Anwar, 2001).

Introductions

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Introduced toIntroduced fromYearReasonIntroduced byEstablished in wild throughReferencesNotes
Natural reproductionContinuous restocking
Colombia Brazil 1960s Seed trade (pathway cause) Yes Millhollon and Burner (1993)
Louisiana Arkansas 1982 Hitchhiker (pathway cause) Yes Arkansas State Plant Board (2008) Introduced by crop debris on farm equipment. Subject to control programme
North Carolina Louisiana 1970s-1980s Hitchhiker (pathway cause) Yes Hall and Patterson (1992) Along railroad
USA Philippines Early 1900s Forage (pathway cause) Yes Bugwood (2008)

Risk of Introduction

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The species is one of the most frequently intercepted of the designated “Federal noxious weeds” found by agricultural inspectors at the USA border, as seeds often hitchhike on railroad cars from Mexico (Lehtonen, 2003). Seed has been found as a contaminant of the following commodities: beans (Phaseolus sp.) from El Salvador and Honduras; false coriander seeds (Eryngium foetidum) from Trinidad; flax seeds (Linum usitatissimum) from El Salvador; mung bean (Vigna radiata) from Belize; salvia seeds (Salvia officinalis) from China; sorghum seeds (Sorghum bicolor) from Costa Rica; sesame seeds (Sesamum indicum) from India and Sudan; and turkeyberry fruit (Solanum torvum) from El Salvador. Continued vigilance is required to prevent further movement in seed samples distributed by research organizations or by seed merchants.

Further movement in soil or plant debris on construction or agricultural machinery is likely as demonstrated by the recent introduction of the species from Louisiana to Arkansas on a combine used in previous harvesting years (Arkansas State Plant Board, 2008).

Habitat

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R. cochinchinensis is a weed of warm-season crops in a variety of habitats around the world. It also grows along roadsides and in other open, well-drained sites. It is an important species in old field succession. However, it also grows in wet places, and even in shallow water. It occurs in habitats with full sun, moderate shade, or even thickets and forests. R. cochinchinensis is most troublesome between 800 and 1300 m in elevation, with rainfall the main limiting factor below 1300 m, and temperature the main limiting factor above (Holm et al., 1977). In eastern Africa, R. cochinchinensis is one of the primary colonizers of disturbed land, and it displaces early perennial colonizers such as Bermuda grass (Cynodon dactylon) and purple nutsedge (Cyperus rotundus) in Trinidad (Holm et al., 1977).

Habitat List

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CategoryHabitatPresenceStatus
Terrestrial-managed
Cultivated / agricultural land Principal habitat Harmful (pest or invasive)
Disturbed areas Principal habitat Harmful (pest or invasive)
Managed forests, plantations and orchards Principal habitat Harmful (pest or invasive)
Rail / roadsides Principal habitat Harmful (pest or invasive)
Terrestrial-natural/semi-natural
Natural forests Secondary/tolerated habitat Natural
Natural forests Secondary/tolerated habitat Productive/non-natural
Natural grasslands Secondary/tolerated habitat Harmful (pest or invasive)
Riverbanks Secondary/tolerated habitat Harmful (pest or invasive)
Riverbanks Secondary/tolerated habitat Natural

Hosts/Species Affected

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R. cochinchinensis is an aggressive weed under various ecological conditions, in at least 18 crops in 44 countries. R. cochinchinensis is also a weed of bananas, cassava, citrus, cowpeas, papayas, groundnut, pineapple, rice, and sorghum in Cuba, Ghana, Jamaica, the Philippines, Trinidad, Venezuela and much of Central America. (Holm et al., 1977; Tabora, 1979; Valverde, 2003; Vibrans, 2009).

Host Plants and Other Plants Affected

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Growth Stages

Top of page Seedling stage, Vegetative growing stage

Biology and Ecology

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Genetics

R. cochinchinensis has evolved distinct biotypes, many of which have been distinguished in different areas of its range. Most of its biotypes were diploids with 2n=20 chromosomes, but some were polyploids (2n=40 or 2n=60). The shape of the glume tip on the pedicellate spikelet was found to be acuminate, acute or obtuse for the biotypes with 2n=20, 40 and 60 chromosomes, respectively, and this was a reliable marker to distinguish diploids from polyploids (Millhollon and Burner, 1993). The receptor of photoperiodic stimulus appears to be the expanding leaf. Alves et al. (2003) also distinguished both diploid and polyploid biotypes from São Paulo State in Brazil, differing in seed size, stomatal size and total chromosome length, variability that was confirmed at the molecular level.

Cytological and morphological studies of two morphotypes of R. cochinchinensis, collected from Kerala State, India, revealed that the short morphotype was diploid (2n=20) and the tall robust morphotype was tetraploid (2n=40). Cytological data indicated that the short morphotype was one of the putative parents of the tetraploid. Apart from variations in the size of the plant and its organs, the two morphotypes differed considerably in the size of sessile spikelets, the size and shape of pedicelled spikelets and the morphological features of the glumes of both spikelets. It is suggested that R. cochinchinensis is an example of a species that has reached the stage of a young polyploid complex (Christopher et al., 1989).

Amplified fragment length polymorphism (AFLP) analysis has indicated an extremely narrow genetic base among accession of the species, possibly as a result of the predominantly inbreeding nature of the weed and to the relatively recent expansion of its geographic range According to Valverde (2003) AFLP studies have identified five major biotype groups closely related to their geographical distribution. Two major groups were comprised of biotypes predominantly collected in Latin America, suggesting that the majority of Latin American biotypes might have arisen from a limited number of introductions, probably from Africa and some from Asia.

Reproductive Biology

Reproduction of R. cochinchinensis is by seeds. The floral structure allows for self-pollination with occasional cross pollination (Mercado, 1978). In the Philippines, the plant flowers all year long, and a single plant may produce 2000-16,000 seeds. Seed production begins 6-7 weeks after emergence and generally continues throughout the growing season. Studies in Zimbabwe have shown that dense stands of the plant will produce over 600 kg of seed per season (Holm et al., 1977). There is no clear correlation of floral response to day length with distribution, but Millhollon and Burner (1993) demonstrated some tendency for African populations to have short-day behaviour. The seeds fall from the plant as they mature, ensuring a continuous supply of new seed on the ground. 

Physiology and Phenology

R. cochinchinensis is regarded as a C4 grass (Das et al., 1993), and growth is very rapid under favourable conditions. Millhollon and Burner (1993) found variation in photoperiodic sensitivity. Whereas all flowered at 12 hours day length, about half were day-neutral and flowered almost as quickly at 14 hours, and some behaved as short-day plants and remained vegetative at 14 hours day length. There was no clear correlation with distribution, but some tendency for African collections to have short-day behaviour, being placed in five broad groups based primarily on the effect of day length on flowering but also on general morphology and pattern of growth (e.g. time until flowering). 

The seeds require a dormancy period of 5-6 months following maturity, with the result that germination occurs in the season following dispersal or in subsequent years. There appear to be two dormancy mechanisms operating in R. cochinchinensis seeds. The major mechanism is imposed by the covering structures and may prevent entry of oxygen into the seed, which stimulates the pentose phosphate pathway. The second mechanism appears to be influenced by light (Clavijo, 1978). Light apparently had an inhibitory effect on germination of in-husk seed but no effect on dehusked seed. In Bolivia, a positive correlation has been shown between rainfall and emergence of R. cochinchinensis. The average depths from which R. cochinchinensis seedlings emerged were 1.6 cm on heavy soil and 5 cm on light soil. In laboratory trials, the optimum temperature for germination was 25°C. Germination was highest in loam soil and lowest in sand soils. The effect of soil moisture content on germination was dependent on soil type, with the moisture required for germination decreasing with increasing tendency of the soil to waterlogging (Unterladstatter, 1979). The seed remains viable in the soil for up to 4 years (Thomas and Allison, 1975), though in southern Benin, it was found that only around 1% of seeds of R. cochinchinensis survived 2 years' burial 3-10 cm deep in soil (Lutzeyer and Kock, 1992). In Zimbabwe, it was found that virtually all seeds of R. cochinchinensis lost their viability within five years (Schwerzel, 1976). 

Environmental Requirements

R. cochinchinensis is reported as a tropical weed mostly from latitude 23°N to 23°S. It also has the ability to grow, flower and set seed under some temperate climate regimes found in the USA in the Gulf Coast states, the lower Midwest, the South Atlantic states and the South-West, where it can reach 75-100% of its growth potential (Patterson et al., 1979). The potential ecological range of R. cochinchinensis in the USA was investigated under controlled conditions in Mississippi (Patterson and Quimby, 1979), and on the basis of its growth responses to temperature, R. cochinchinensis represents a serious potential weed problem in these regions. Controlled field studies as well as other laboratory experiments have shown that R. cochinchinensis can grow and produce seed as far north as St. Paul, Minnesota, USA (Millhollon, 1975).

Climate

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ClimateStatusDescriptionRemark
Am - Tropical monsoon climate Preferred Tropical monsoon climate ( < 60mm precipitation driest month but > (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])
BSh - Steppe climate Tolerated > 430mm and < 860mm annual precipitation, low altitude, average temp. > 18°C
Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year
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)

Latitude/Altitude Ranges

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Latitude North (°N)Latitude South (°S)Altitude Lower (m)Altitude Upper (m)
17 25

Air Temperature

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Parameter Lower limit Upper limit
Mean annual temperature (ºC) 19 30
Mean maximum temperature of hottest month (ºC) 22 32
Mean minimum temperature of coldest month (ºC) 5 26

Rainfall

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ParameterLower limitUpper limitDescription
Dry season duration07number of consecutive months with <40 mm rainfall
Mean annual rainfall6303590mm; lower/upper limits

Rainfall Regime

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Summer

Soil Tolerances

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Soil drainage

  • free

Soil reaction

  • acid

Soil texture

  • heavy
  • medium

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Colletotrichum sp. near graminicola Pathogen Leaves
Gibberella fujikuroi Pathogen
Puccinia rottboelliae Pathogen Leaves
rottboellia yellow mottle virus Pathogen
Sporisorium ophiuri Pathogen Leaves to species
Steneotarsonemus furcatus Herbivore Leaves not specific

Notes on Natural Enemies

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Investigations of natural enemies of R. cochinchinensis have concentrated on fungal pathogens, as these have been considered to have potential for use in biocontrol programmes in Central America. The most studied is the head smut Sporisorium ophiuri, which has a widespread distribution in Africa and Asia (Ellison, 1987, 1993; Reeder et al., 1996). The smut is a soil-borne, systemic pathogen, infecting itchgrass seedlings before they emerge from the soil. In the natural range of R. cochinchinensis, natural epiphytotics of the smut are common, often infecting a high percentage of plants within a population. Other pathogens associated with the grass in the Old World Tropics include a Colletotrichum sp. near graminicola from Thailand (Ellison and Evans, 1995) and the leaf rust Puccinia rottboelliae (Ellison and Bird, 1996). Sanchez Garita and Zuniga (1999) reviewed the occurrence of indigenous pathogens on R. cochinchinensis in Latin America and reported species from the genera Cladosporium, Curvularia, Drechslera, Fusarium, Helminthosporium and Pestolita, and two bacteria, Pseudomonas and Xanthomonas were also reported to occur on the weed.

Means of Movement and Dispersal

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Natural Dispersal (Non-Biotic)

Seeds are spread by water (Holm et al., 1977) and flood water (Freshwater et al., 1986). 

Vector Transmission (Biotic)

Seed can be spread from infested areas to non-infested areas by wild and domesticated birds (Aison et al., 1984). In Zimbabwe (Thomas and Allison, 1974), 2% of R. cochinchinensis seeds fed to leghorn hens were recovered from the droppings and most of them were found to be viable. Faeces of the slender mongoose (Herpestes sanguineus) were also found to contain many R. cochinchinensis seeds. The results suggest that only birds or animals producing faeces that are fairly coarse or contain some roughage are likely to disseminate R. cochinchinensis, and then only if the seeds forms part of their diet (Thomas and Allison, 1974). 

Intentional Introduction

There is anecdotal evidence for the intentional introduction of R. cochinchinensis as a potential pasture species into the southern USA in the early 1900s (Bugwood, 2008), and R. cochinchinensis was first introduced into the USA from the Philippines in the early 1900s as a potential pasture grass (Bugwood, 2008). However although the species can produce high yields as a forage it is not as palatable as other options (Bwire et al., 2003) so is unlikely to be the subject of further deliberate introduction. 

Accidental Introduction:

Contamination of samples of crop seed germplasm for research, of commercial seed lots and commodities has provided a common route for accidental introduction of R. cochinchinensis in Latin America and southern USA (e.g. Millhollon and Burner, 1993; Lehtonen, 2003). At a regional and local level, subsequent introductions of seed in soil or plant debris on construction or agricultural machinery has been well documented (Millhollon, 1980; Arkansas State Plant Board, 2008). Other introductions into southern USA early in the 1900s are thought to have come via the West Indies in plant material or on equipment (Millhollon, 1965). The spread of R. cochinchinensis in Louisiana has been attributed in part to movement of equipment and road matting materials used during the extensive oil exploration that occurred in the late 1970s and early 1980s. Machinery was also responsible for the introduction of the species into Arkansas from seed carried in crop debris on a combine harvester used in previous harvest in Louisiana (Arkansas State Plant Board, 2008). Its appearance in south-eastern North Carolina in the early 1980s along the north-south seaboard coastline railroad is an example of this mode of spread (Hall and Patterson, 1992), and it is one of the most frequently intercepted Federal noxious weeds found by Agricultural inspectors at the USA border as seeds often hitchhike on railroad cars from Mexico (Lehtonen, 2003). Vibrans (2009) reports that due to dispersal on machinery, invasion often starts at roadsides.

Introductions have also been through contaminated crop seed for trials exchanged by international research programmes or in commercial seed supplied to farmers. For example, Rottboellia has been found in rice seed lots received at the International Rice Research Institute in the Philippines (Huelma et al., 1996). Similarly, there are indications of dissemination in rice seed movements from Colombia to Brazil in 1961 (Millhollon and Burner, 1993). In Campeche and other areas in Mexico, farmers identified contaminated rice seed as being responsible for the introduction of R. cochinchinensis to their fields (Valverde et al., 1999b, 2001). Movement of livestock and farm machinery is blamed for the introduction of an exotic biotype from Thailand to Malaysia (Anwar, 2001).

Pathway Causes

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CauseNotesLong DistanceLocalReferences
Breeding and propagationE.g. in contaminated rice for research purposes at IRRI Yes Yes Huelma et al., 1996
Crop productionContaminated farm machinery Yes Arkansas State Plant Board, 2008
Digestion and excretion Yes Thomas and Allison, 1974
Flooding and other natural disasters Yes Freshwater et al., 1986
ForageOriginally to the USA, but unlikely now Yes Bugwood, 2008
HitchhikerRailcars from Mexico to USA Yes Lehtonen, 2003
ResearchItchgrass seed found as contaminant of seed in germplasm collections received at IRRI Yes Yes Huelma et al., 1996; Murphy and Cheesman, 2006
Seed tradeContaminated rice seed source of introduction in Campeche, Mexico Yes Yes Valverde et al., 1999; Valverde et al., 2001
StockingThailand to Malaysia with livestock Yes Yes Anwar, 2001

Pathway Vectors

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VectorNotesLong DistanceLocalReferences
Bulk freight or cargoContaminated grain Yes Valverde et al., 1999; Valverde et al., 2001
Containers and packaging - wood3 litre pots Yes
Land vehiclesRail trucks from Mexico to USA Yes Lehtonen, 2003
Machinery and equipmentThailand to Malaysia, and in southern USA Yes Yes Anwar, 2001; Arkansas State Plant Board, 2008; Millhollon, 1980
Plants or parts of plantsAs a traded seed contaminant Yes Millhollon and Burner, 1993
Soil, sand and gravelRoad construction aggregate in Louisiana Yes Millhollon, 1980
Water Yes Holm et al., 1977

Plant Trade

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Plant parts liable to carry the pest in trade/transportPest stagesBorne internallyBorne externallyVisibility of pest or symptoms
True seeds (inc. grain)
Plant parts not known to carry the pest in trade/transport
Flowers/Inflorescences/Cones/Calyx
Stems (above ground)/Shoots/Trunks/Branches

Impact Summary

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CategoryImpact
Cultural/amenity Negative
Economic/livelihood Negative
Environment (generally) Negative
Human health Negative

Economic Impact

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R. cochinchinensis is an aggressive weed under various ecological conditions, in at least 18 crops in 44 countries. R. cochinchinensis is a serious weed of cotton in Zambia and Zimbabwe and a moderate weed of cotton in Ethiopia, Mozambique, Sudan, Uganda and Venezuela; a serious weed of groundnut in Sudan, Zimbabwe and Zambia, and a moderate weed of groundnut in Trinidad; a serious weed of mungbean in the Philippines, and a moderate weed of sesame in India. R. cochinchinensis is also a weed of bananas, cassava, citrus, cowpeas, papayas, groundnut, pineapple, rice, and sorghum in Cuba, Ghana, Jamaica, the Philippines, Trinidad and Venezuela (Holm et al., 1977; Tabora, 1979). In addition to its effects on crop yield, R. cochinchinensis is a problem to labourers, as the needle-like hairs on the leaf sheaths break off in the skin and can cause painful infections.

R. cochinchinensis is a serious weed of soyabean in the Philippines, Zambia and Zimbabwe. Soyabean seed weight within 20 cm of the weed was reduced by 15-21% and a seed weight reduction of 9% was detected at a distance of 40-60 cm from the weed. Weight reductions were attributed to decreases in seed number of 12-22% within 40 cm of the weed. R. cochinchinensis interference increased soyabean height within 40 cm of the weed, but soyabean canopy width was generally unaffected.

R. cochinchinensis is a serious weed of sugarcane in the Philippines, USA and Zambia, and a moderate weed of sugarcane in Kenya, Madagascar and Venezuela. R. cochinchinensis removed at harvest (after 180 days of interference) reduced sugar yield by 19% in four experiments. R. cochinchinensis infestations similarly removed in the second-year crop reduced sugar yield by a maximum of 72% compared to a 2-year weed-free control. R. cochinchinensis interference reduced sugar yield primarily by reducing stalk population even though full-season interference increased sugar concentration of juice by 2-10%. These studies indicate that R. cochinchinensis must be removed from sugarcane well before 30 days of interference under Louisiana growing conditions. However, sugarcane stands and yield recovered almost completely when maintained weed-free in the second-year crop following full-season R. cochinchinensis interference in the first-year crop (Millhollon, 1992).

R. cochinchinensis is a serious weed of rice in the Philippines. Uncontrolled weeds dominated by R. cochinchinensis decreased yields of rice cv. Kinandang Pula to 50 kg/ha, whereas a single weeding increased yields to about 2000 kg/ha and double weeding to about 2500 kg/ha (Legaspi et al., 1989). However, no significant difference was found between times of weeding. Complete weed control produced 3000 kg/ha of grain.

R. cochinchinensis is a serious weed of maize in Ghana, Philippines, Zambia and Zimbabwe, and a moderate weed of maize in Colombia, Nigeria, Tanzania and Venezuela. Studies have shown that an R. cochinchinensis density of 50 plants/m² can reduce maize yields by almost 50%. A density of 142 plants/m² can cause reductions up to 71% (Mercado, 1978). In the maize-growing sectors of Mindanao, Philippines, R. cochinchinensis has been recognized as the main weed menace. Uncontrolled R. cochinchinensis growth reduced yields by 63 to 71%. Results from hand-weeding experimental plots revealed that local farmers suffer a 24% average crop loss from this weed alone when the normal procedure of one mechanical cultivation and a hilling operation are followed. This cultivation misses weeds in the crop row and hand-weeding is not a common practice. Also, R. cochinchinensis competes throughout the season and can exert its most damaging effect late in the season. Monocrop maize and the associated cultural practices tend to encourage weed growth. Also, regional maize-growing farms generally fall in the 3-5 ha range, thus precluding sufficient time or labour to weed the planted area adequately.

In addition to herbicide treatments, two other approaches for control were investigated. Adding one correctly timed hand-weeding to the present mechanical cultivation-hilling routine could cut weed-caused yield losses to approximately 5%. Growing mungbean (Vigna radiata) between the maize rows not only reduced the weed competition but also provided additional income as mungbean sells at 6-10 times the value of maize (NCPC, 1979). In Costa Rica, control costs in maize might represent up to 26 percent of the income obtained from selling the grain (Valverde et al., 1999b).

R. cochinchinensis is also an alternative host of the viruses causing corn leaf gall and rice leaf gall (Agati and Calica, 1949). R. cochinchinensis has been reported as an alternative host of Oligonychus grypus, a pest of sugarcane (Gutierez and Gonzalez, 1993). It has also been reported as a host of Diopsis [Diasemopsis] macrophthalma, an insect pest of rice (Alghali and Domingo, 1982). Curvularia cymbopogonis, which is a pathogen of rice in the Philippines, was isolated from diseased R. cochinchinensis plants collected in southern Louisiana in the late 1970s. The disease was subsequently found to be pathogenic to seedlings of the weed grown in a growth chamber (Walker and White, 1979). In 1980, it was reported that R. cochinchinensis was found to be an alternative host of a virus-like disease of maize in Guadeloupe, French West Indies. The causal agent was found to be transmitted in a persistent manner by Peregrinus maidis. An isolate of Hoja blanca maize virus from Venezuela and an extract of a sample of R. cochinchinensis from Guadeloupe proved positive against Maize stripe virus from Florida (Miglioria and Lastra, 1980).

Environmental Impact

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Studies of this species have focused on impacts in crops rather than on environmental impacts. R. cochinchinensis has become a common component of roadside vegetation and on other disturbed areas in Central America but no information is available on impacts on biodiversity and there are no reports of threats to the native flora. In Costa Rica and Cuba this species is considered a habitat transformer and it is listed among the 100’s most noxious invasive plants in these countries (Chacon and Saborio, 2012; Oviedo Prieto et al., 2012).

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
  • Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
  • Pioneering in disturbed areas
  • Highly mobile locally
  • Fast growing
  • Has high reproductive potential
  • Has propagules that can remain viable for more than one year
  • Has high genetic variability
Impact outcomes
  • Damaged ecosystem services
  • Host damage
  • Modification of successional patterns
  • Monoculture formation
  • Negatively impacts agriculture
  • Negatively impacts human health
  • Negatively impacts livelihoods
  • Reduced amenity values
  • Threat to/ loss of native species
Impact mechanisms
  • Causes allergic responses
  • Competition - monopolizing resources
  • Competition - shading
  • Pest and disease transmission
  • Parasitism (incl. parasitoid)
  • 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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R. cochinchinensis is not generally considered to have much economic value. It has been tested as a fodder grass and this is said to be how it was first introduced into USA in the early 1900s (Bugwood, 2008), however, it has rarely been exploited as a forage in its native range.

Uses List

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Animal feed, fodder, forage

  • Forage

Similarities to Other Species/Conditions

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R. cochinchinensis superficially resembles johnsongrass (Sorghum halepense) and other tall weedy grasses. However, it can be distinguished from other grasses by the trichomes (stiff hairs) present on the leaf sheaths responsible for the common name itch grass. Another diagnostic feature is the jointed cylindrical seed head. The seedheads (spikes) are produced on the ends of the ends of the main shoots as well as on axillary branches and tiller shoots.

Mnesithea selloana(syn. Rottboellia selloana) is a tufted perennial grass growing to 30-65 cm high with leaves 12-20 cm long, found in Uruguay, southern Brazil and north-eastern Argentina in dry grasslands on sandy soil, being well grazed by cattle (Bogdan, 1977). The inflorescence is similar to R. cochinchinensis but individual seeds are somewhat smaller.

Prevention and Control

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SPS Measures

R. cochinchinensis  is listed as a noxious weed under USA Federal law (Federal Noxious Weed List), and is also prohibited in Arkansas, Mississippi, Florida, Georgia, and North and South Carolina. Observation of seed during inspection of commodity shipments e.g. border control from Mexico to USA can help prevent introduction (Lehtonen, 2003). 

Control

Physical/mechanicalcontrol

Cultural control is based on exhausting the seed bank. The effect of cultivation every 2, 3 or 5 weeks on R. cochinchinensis emergence in a fallow field in the Philippines, after 1 year, substantially decreased the R. cochinchinensis population; and the greater the frequency of cultivation, the greater the population decrease. By the end of the second year, R. cochinchinensis had been practically eliminated by cultivation every 2-3 weeks, and only a few plants grew on plots cultivated every 5 weeks. Total weed density after 2 years was not reduced in plots rotary cultivated every 5 weeks, but R. cochinchinensis was replaced by less competitive weeds (Navarez et al., 1987). Mechanical weeding with a tool that throws the earth and weeds to the row centre (‘off-barring’) and earthing up, were found to be inadequate for control of R. cochinchinensis in maize in the Philippines, but supplementary weedings, either manual or chemical, substantially increased both yields and net profits (Pamplona, 1981). Uncontrolled R. cochinchinensis reduced maize yields by about 50%, whereas two cultivations using draft animals controlled R. cochinchinensis to some extent but yields were still reduced by 24%, and traditional hand-weeding three times failed to reduce the R. cochinchinensis infestation population in the maize row. The best control of R. cochinchinensis was achieved by two mechanical cultivations plus hand-weeding in the maize row, and the best economic return was achieved by inter-cropping maize with mung bean (Fisher et al., 1980). 

Reduction of itchgrass infestation in roadside vegetation has been achieved in south-western Louisiana by careful manipulation of the annual mowing frequency (Venner, 2006). 

Biological control

Isolates of fungi collected from R. cochinchinensis have been screened for host specificity and three of them selected for further study as potential biological control agents. An isolate of Colletotrichum sp. near graminicola from Thailand was tested in the laboratory and in field trials in Thailand as a possible candidate for development as a mycoherbicide. The results were equivocal but a synergistic response was found when a low dose of chemical herbicide was added to the fungal inoculum (Ellison and Evans, 1995). As the sporidia are readily produced in culture, it may be possible to apply them to the soil as a form of mycoherbicide or introduce the fungus as a classical biocontrol agent (Ellison and Evans, 1995). Isolates of the smut are itchgrass-biotype specific but one from Madagascar was found to infect a wide range of biotypes including a number from Latin America, and hence selected for a comprehensive host range screening. The smut was found to be extremely host specific, and none of 49 species/varieties of graminaceous test plants other than itchgrass became infected. Screened species included pastures, weedy grasses, graminaceous crops (rice, sugar cane, maize, sorghum) and teosinte (Zea (Euchlaena) mexicana), the maize ancestor (Reeder and Ellison, 1999). The potential efficacy of this pathogen as a classical agent lies in the short-lived nature of the weed-seed bank (three to four years) and the aggressiveness of the smut. The Costa Rican plant health authorities approved the introduction of the smut in December 1999 into quarantine for additional host range screening, and it was hoped to eventually attract funding for field release (Ellison and Barreto, 2004). 

A rust, Puccinia rottboelliae, causes severe seedling infection in the field and preliminary host-range tests with an isolate from Kenya suggest that it is specific to R. cochinchinensis. Thus, this rust may have potential as a classical biological control agent in the Americas, perhaps involving a management strategy including early-season augmentation, and further research is being carried out on these agents in order to develop a biological control strategy (Ellison and Bird, 1996).

The fungal pathogen Exserohilum longirostratum has been investigated in Malaysia as a potential biocontrol agent. Kadir et al. (2007) reported that applied as a post-emergence foliar spray, the fungus inflicted high percentage of mortality to young itch grass seedlings. It did not kill older plants but was capable of reducing biomass by approximately 56%. Patel and Patel (2014) also found in India that this fungus was highly pathogenic on itch grass. Alloub et al. (2009) suggest that Exserohilum prolatum [Setosphaeria prolata] has good potential for biocontrol, especially when applied multiple times.  

Chemical control

Chemical control of R. cochinchinensis is made difficult in many crops by the resistance of this species to many herbicides in important groups such as the triazines (e.g. atrazine etc.) and chloroacetanilides and to others such as metribuzin and terbacil. Populations of R. cochinchinensis that have evolved resistance to herbicides that inhibit the enzyme acetyl coenzyme-A carboxylase have been identified in Bolivia (Avila et al., 2007), including haloxyfop-R-methyl and sethoxydim.

In sugarcane plantations in Jamaica, mixtures containing asulam, 2,4-D + ioxynil and the cationic surfactant ethokem (polyethanoxy amine) effectively controlled R. cochinchinensis (and several other grasses). The inclusion of ethokem in the mixture provided excellent control of R. cochinchinensis, and enabled a reduction in the amount of other ingredients (Anon., 1993). In Louisiana, nicosulfuron, primisulfuron, paraquat and ametryn treatments were compared to the asulam standard for post-emergence treatment. Nicosulfuron provided the greatest season-long control of R. cochinchinensis. Asulam and nicosulfuron did not reduce late-season R. cochinchinensis infestation levels when compared to the control, whereas post-emergence applications of paraquat and ametryn reduced late-season infestations by 74 and 52%, respectively. It was concluded that paraquat and ametryn applied post-emergence provided R. cochinchinensis control without reducing sugarcane yields (Lencse et al, 1992). Also, R. cochinchinensis control and sugarcane injury with both soil-incorporated and soil surface-applied herbicide treatments were compared with the standard pre-emergence trifluralin treatment. The soil-incorporated treatments consisted of pendimethalin, fomesafen and clomazone. Results were comparable to those with soil-incorporated trifluralin. The soil-surface herbicide applications (pendimethalin + atrazine, prodiamine, clomazone and fomesafen) provided acceptable (>80%) R. cochinchinensis control when compared to untreated sites. Fomesafen, quinclorac, terbacil, metribuzin and atrazine provided poor R. cochinchinensis control. Sugarcane injury was minimal, with the exception of that caused by clomazone, which caused a temporary bleaching of sugarcane foliage. It was concluded that, compared with soil-incorporated trifluralin, the use of pendimethalin, prodiamine, fomesafen and clomazone applied to the soil surface would reduce trips across the field and may minimize sugarcane injury, but may not provide consistent R. cochinchinensis control when rainfall is not received for activation (Griffin and Lencse, 1992). R. cochinchinensis control was 86% with non-incorporated pendimethalin and prodiamine. Standard incorporated trifluralin resulted in 99% R. cochinchinensis control. It is concluded that, when used in conjunction with other herbicides, non-incorporated pendimethalin and prodiamine are effective alternatives to soil-incorporated treatments for weed control in sugarcane (Millhollon, 1993). 

Weed control measures in a rice-maize-soyabean rotation in Peru identified weed species resistant to the herbicide programme in a continuous cropping system, and effective weed management practices for intensively managed cropping systems in the humid tropics were developed. Composition of first-crop weeds was estimated to be 60% grass, 25% sedges, and 15% broadleaved weeds. R. cochinchinensis comprised 85% of the grass infestation. Metolachlor controlled other grasses and most broadleaf weeds in maize and soyabean but it did not control R. cochinchinensis. Metolachlor alone resulted in a weed population that was 97% R. cochinchinensis in the sixth crop. Sethoxydim + bentazone on soyabean controlled grasses including R. cochinchinensis, but broadleaved weeds increased. Propanil + oxadiazon on rice resulted in a mixed grass population. Rice was more vulnerable to weed pressure than either maize or soyabean and appears inappropriate for this high-input rotation because of the high cost of weed control (Pleasant et al., 1990). 

In field trials in soyabeans, trifluralin and pendimethalin incorporated pre-sowing gave 87 and 78% control of R. cochinchinensis, respectively, in late season when averaged over a 2-year period. Clomazone applied pre-emergence to the soil surface in one year controlled 80% of R. cochinchinensis compared with 51% when incorporated pre-sowing. Post-emergence application of fluazifop-P, haloxyfop, quizalofop and diclofop gave at least 90% control. R. cochinchinensis was less well controlled by sethoxydim and clethodim post-emergence, while imazaquin and imazethapyr lacked adequate activity (Griffin, 1991). In tests in southern Louisiana, overall application of diclofop gave 73-94% control of 10-30 cm tall R. cochinchinensis. Diclofop was effective as a follow-up, mid-season treatment whereas dinitroaniline was used pre-sowing and this combination resulted in higher yield increases than those obtained with mid- or late-season applications of diclofop alone (Nester and Harger, 1980). In an earlier study conducted in Louisiana, USA, dinitroaniline herbicides which showed 80% or more control of R. cochinchinensis in soyabeans were (1) on fine-textured soil: trifluralin, fluchloralin and pendimethalin, prodiamine and profluralin, and (2) on medium-textured soil: trifluralin, profluralin and pendimethalin, prodiamine and fluchloralin (Vidrine et al., 1979).

In Nigeria, metolachlor, chlorthal-dimethyl and pendimethalin applied as pre-emergence controlled most annual grasses (including R. cochinchinensis) in cowpea (Vigna unguiculata). None of these herbicides injured the crop so that cowpea yields from the herbicide treatments were as good as those from normal hand-weeding or experimental clean-weeding (Akobundu, 1982). 

Treatments controlling R. cochinchinensis in maize in the Philippines giving yields comparable to a hand-weeded control were: off-barring (moving soil away from the crop row into the inter-row), and hand weeding in the row at 12 days after sowing followed by earthing-up at 26 days after sowing band application of trifluralin between the rows, followed by either hand-weeding in the rows at 12 days after sowing, earthing-up at 26 days after sowing, or both; off-barring and hand weeding in the rows at 12 days after sowing followed by paraquat/2,4-D application; and pre-emergence application of atrazine, in combination with either earthing-up or directed paraquat/2,4-D application at 26 days after sowing (Pamplona and Imlan, 1977). The ALS inhibitor nicosulfuron, applied with a nonionic surfactant also provides effective control when applied alone to R. cochinchinensis at the 6 leaf stage (Strahan et al., 2000). Work on herbicide mixtures and sequences in Louisiana demonstrated that itchgrass was controlled 71% when imazethapyr plus imazapyr was applied alone post-emergence, but control was nearly 86% when pendimethalin or nicosulfuron was applied with the imazethapyr plus imazapyr mixture (Bond and Griffin, 2005). 

Studies in Venezuela for controlling heavy grass weed infestations in potatoes recommended using metribuzin. The most effective product for control of R. cochinchinensis was found to be pendimethalin (Anon., 1977). Labrada (1994) provides a useful summary of herbicide use in a range of vegetable and other crops, based on trifluralin, pendimethalin, oxadiazon, clomazone, diphenamid, napropamide, fluazifop, haloxyfop and quizalofop. 

IPM programmes

In East Africa, R. cochinchinensis is controlled by a combination of cultivation followed by fallowing for at least 2 years. The infested site is first burned to destroy the seeds on the surface. Next, it is ploughed to stimulate germination of seeds in the top soil horizon. Following this, deep ploughing is done to bury the seedlings. After this, the land is left fallow until the buried seeds expire and the land is considered clean (Holm et al., 1977). Preventing seed production is the most effective method of controlling this species (Schwerzel, 1976; Lutzeyer and Kock, 1992). 

In field trials conducted in Costa Rica, the effects of manual weed control, disc ploughing, pendimethalin and paraquat were evaluated in different combinations for the control of R. cochinchinensis in maize-bean [Phaseolus spp.] rotations. All weed control treatments reduced R. cochinchinensis plant density from untreated controls. Pendimethalin resulted in the greatest weed control in the crop (Rojas et al., 1993). Itchgrass density was substantially higher in plots without control in the fallow period but use of in-crop herbicides decreased the weed populations to similar levels, regardless of fallow management. Lower itchgrass populations also were observed in plots with zero tillage compared with conventional tillage. R. cochinchinensis densities were also reduced by integrating no-tillage, use of the selective herbicide pendimethalin in the first maize crop (to lower the initial density of itchgrass), planting of a velvetbean cover crop between maize rows, and prevention of itchgrass seed set in the fallow period (Valverde et al., 1999a, b).

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Links to Websites

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WebsiteURLComment
Invasive Species Specialist Group (ISSG) website on T. melanocephalumhttp://www.issg.org
North American Plant Protection Organizationhttp://www.pestalert.org

Contributors

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01/07/14 Updated by:

Julissa Rojas-Sandoval, Department of Botany-Smithsonian NMNH, Washington DC, USA

Pedro Acevedo-Rodríguez, Department of Botany-Smithsonian NMNH, Washington DC, USA

06/06/2008 Updated by:

Charlie Riches, Consultant, UK

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