Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide


Setaria viridis
(green foxtail)



Setaria viridis (green foxtail)


  • Last modified
  • 22 November 2019
  • Datasheet Type(s)
  • Invasive Species
  • Pest
  • Host Plant
  • Preferred Scientific Name
  • Setaria viridis
  • Preferred Common Name
  • green foxtail
  • Taxonomic Tree
  • Domain: Eukaryota
  •   Kingdom: Plantae
  •     Phylum: Spermatophyta
  •       Subphylum: Angiospermae
  •         Class: Monocotyledonae

Don't need the entire report?

Generate a print friendly version containing only the sections you need.

Generate report


Top of page
S. viridis growing in the USA, showing inflorescence (usually 15 x 1 cm, dense spike-like panicle, erect or slightly nodding at the tip).
TitleMature plants
CaptionS. viridis growing in the USA, showing inflorescence (usually 15 x 1 cm, dense spike-like panicle, erect or slightly nodding at the tip).
Copyright©Chris Parker/Bristol, UK
S. viridis growing in the USA, showing inflorescence (usually 15 x 1 cm, dense spike-like panicle, erect or slightly nodding at the tip).
Mature plantsS. viridis growing in the USA, showing inflorescence (usually 15 x 1 cm, dense spike-like panicle, erect or slightly nodding at the tip).©Chris Parker/Bristol, UK


Top of page

Preferred Scientific Name

  • Setaria viridis (L.) Beauv. (1812)

Preferred Common Name

  • green foxtail

Other Scientific Names

  • Chaetochloa viridis (L.) Scribn.
  • Chamaeraphis viridis (L.) Millsp. (1892)
  • Ixophorus viridis (L.) Nash (1895)
  • Panicum bicolor Moench. (1794)
  • Panicum laevigatum Lam.
  • Panicum purpurascens Opiz (1823)
  • Panicum reclinatum Vill. (1778)
  • Panicum viride L. (1759)
  • Pennisetum viride (L.) R. Br. (1822)

International Common Names

  • English: bottle grass (Canada); giant green foxtail (USA); green bristlegrass; green panicum; green pigeongrass (Australia); robust purple foxtail (USA); robust white foxtail (USA)
  • Spanish: almoralejo; almorejo verde
  • French: setaire verte
  • Portuguese: capim-verde; milha-verde

Local Common Names

  • Argentina: gramilla
  • Bangladesh: shabuz shiallaja
  • Egypt: deil-el-far
  • Germany: Gròne Borstenhirse; Grònes Fennichgras
  • Iran: arzan
  • Iraq: dukhain el-forsheh
  • Italy: panico selvatico
  • Japan: enokorogusa
  • Netherlands: groene naaldaar
  • Philippines: buntot-pusa
  • Sweden: groen kolvhirs; grønhirs
  • Taiwan: gou-wei-tsau
  • Yugoslavia (Serbia and Montenegro): muraika

EPPO code

  • SETVI (Setaria viridis)

Taxonomic Tree

Top of page
  • Domain: Eukaryota
  •     Kingdom: Plantae
  •         Phylum: Spermatophyta
  •             Subphylum: Angiospermae
  •                 Class: Monocotyledonae
  •                     Order: Cyperales
  •                         Family: Poaceae
  •                             Genus: Setaria
  •                                 Species: Setaria viridis

Notes on Taxonomy and Nomenclature

Top of page
Although this species has many synonyms, Setaria viridis is now accepted as the preferred scientific name. The epithet viridis and the common name, green foxtail refer to the green bristles of the inflorescence. S. viridis is closely related to the cultivated species S. italica (L.) P. Beauv. and is thought to have been its wild progenitor (Wang et al., 1995, 1998). The two species have very similar genomes and are capable of forming hybrids in the wild.

There are a number of varieties or subspecies of S. viridis, some of which have been given their own common names in North America (Douglas et al., 1985). These include: S. viridis var. major (Gaud.) Posp. (giant green foxtail) which was first recognized in North America in 1938 and became common in Illinois and Iowa; var. robusta-alba Schreiber (robust white foxtail); var. robusta-purpurea Schreiber (robust purple foxtail); var. weinmanni (R. & S.) Brand; var. gigantea Fr. et Sav. ex Matsum; forma arenosa (L.) P. Beauv; subsp. glareosa (L.) P. Beauv. and subsp. minor (L.) P. Beauv. Differences in morphology and stature are discussed in the Morphology section.

According to Clayton (1980), hybrids with S. verticillata have been reported, for example, through much of South and Central Europe. However, Stace (1991) concludes that some of these supposed hybrids should be ascribed to S. verticillata var. ambigua.


Top of page
Typical S. viridis (var. viridis) is a tufted annual grass, with many culms, more-or-less erect, up to 70 cm (rarely 100 cm) high. The leaves are about 20 cm (2-40 cm) long by 10 mm (4-25 cm) wide, flat, acuminate, light green, drooping, distinctly, but finely veined with prominent mid-vein below, scabrous above, usually glabrous below. Sheaths are slightly compressed, sometimes purplish at the base, the margins noticeably ciliate. Ligule a fringe of hairs up to 2 mm long, fused at the base. Inflorescence is a dense spike-like panicle, erect or slightly nodding at the tip, up to 15 cm long, about 1 cm in diameter, the rachis often pilose. Spikelets are in very short panicle branches, each spikelet elliptical, up to 2.5 x 1.5 mm wide, subtended by one to three bristles 5-10 mm long, these are usually green, rarely purple, antrorsely barbed (i.e. barbs directed towards the apex, so not tending to stick to clothing as in S. verticillata). The lower glume is one third the length of the spikelet, upper glume 5-6-nerved, almost as long as the lemmas. Lower lemma sterile, like the upper glume, upper lemma fertile, finely rugose. Mature spikelets fall entire, leaving the bristles only (as opposed to S. italica in which the upper fertile floret falls leaving glumes and lower lemma as well as bristles). Chromosome number (2n) = 18. This description is largely based on Douglas et al. (1985). Holm et al. (1977) also provide a description and excellent line drawings, including the seedling stage, showing the ciliate leaf sheath and virtually glabrous leaf blade (unlike S. pumila, which has some long hairs on the upper leaf surface, and no cilia on the sheath).

The development of the root system has been studied and described in some detail (see Douglas et al., 1985).

S. viridis var. major is similar in form to the typical S. viridis var. viridis but much more robust, up to 2 m high with up to 12 nodes per stem (v. 6-7), long nodding inflorescences, brownish red bristles and up to 6000 seeds per panicle (v. 600-800). It has been suggested that this may be a form of S. italica with genes from S. viridis for disarticulation below the glumes (Douglas et al., 1985).

S. viridis var. robusta alba and var. robusta-purpurea differ from typical S. viridis in their much greater vigour and long nodding inflorescence, normally at least 15 cm long, with white and reddish-purple bristles, respectively. They differ from S. viridis var. major in bristle colour and in the denser inflorescence. Numerical and chemotaxonomic studies by Williams and Schreiber (1976) suggest a close relationship with var. major. Schreiber and Oliver (1971) provide a useful key.

S. viridis var. weinmanni has a more spreading habit, narrower leaves and smaller, more slender panicles.


Top of page
S. viridis is a native of Europe but spread to North America as early as 1821 (Douglas et al., 1985) and now occurs in most temperate countries of the Northern and Southern hemispheres. It rarely occurs in the tropics other than at high altitude.

Although listed by Holm et al. (1979) as occurring in Kenya, S. viridis is not recorded by Clayton and Renvoize (1982) for any country in East Africa.

In China, Wang (1980) records S. viridis as occurring 'over all parts of the country'.

In Europe, Clayton (1980) records S. viridis as occurring throughout Europe except the Azores (Portugal), the UK, the Faeroe Islands, Ireland, Iceland and northern Russia. It has, however, been recorded sporadically in the UK.

Distribution Table

Top of page

The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.

Last updated: 25 Feb 2021
Continent/Country/Region Distribution Last Reported Origin First Reported Invasive Reference Notes


EgyptPresent, Widespread
MoroccoPresent, Widespread
NigerPresent, LocalizedNative
South AfricaPresent


ChinaPresent, Widespread
IndiaPresent, Widespread
-Jammu and KashmirPresent, Widespread
-West BengalPresent
IranPresent, Widespread
IraqPresent, Widespread
JapanPresent, Widespread
-Ryukyu IslandsPresent
LebanonPresent, Widespread
MongoliaPresentOriginal citation: Tselev and Federov (1983)
Saudi ArabiaPresent, Widespread
South KoreaPresent
TaiwanPresent, Widespread
TurkeyPresentOriginal recorded location: Turkey-in-Asia; Original citation: Erik and Demirkus, 1988


Federal Republic of YugoslaviaPresent, Widespread
DenmarkPresent, Localized
RomaniaPresent, Widespread
RussiaPresent, Widespread
SpainPresent, Widespread
United KingdomPresent, Localized

North America

CanadaPresent, Widespread
-AlbertaPresent, Widespread
-British ColumbiaPresent, Widespread
-ManitobaPresent, Widespread
-New BrunswickPresent, Widespread
-Newfoundland and LabradorPresent, Localized
-Nova ScotiaPresent, Widespread
-OntarioPresent, Widespread
-Prince Edward IslandPresent, Widespread
-QuebecPresent, Widespread
-SaskatchewanPresent, WidespreadOriginal citation: Douglas et al., 1979
United StatesPresent, Widespread
-New HampshirePresent
-New JerseyPresent
-New MexicoPresent
-New YorkPresent
-North CarolinaPresent
-North DakotaPresent
-Rhode IslandPresent
-South CarolinaPresent
-South DakotaPresent
-West VirginiaPresent


-New South WalesPresent
-Northern TerritoryPresent
-South AustraliaPresent
-Western AustraliaPresent
New ZealandPresent

South America

ArgentinaPresent, Widespread
-GoiasPresentOriginal citation: Flora do Brasil 2020 (2017)
-Minas GeraisPresentOriginal citation: Flora do Brasil 2020 (2017)
-Rio Grande do NortePresent
-Rio Grande do SulPresent
-Sao PauloPresentOriginal citation: Flora do Brasil 2020 (2017)


Top of page
S. viridis is primarily a weed of the temperate zone, and is rarely present in the tropics, other than at high altitudes. It grows mainly in cultivated fields and gardens, but also in waste places, disturbed areas and along roads (Holm et al., 1977).

Habitat List

Top of page

Hosts/Species Affected

Top of page
In many countries, S. viridis is one of the most abundant of all weeds, and it inevitably occurs in most of the crops in those countries, both temperate and sub-tropical. In addition to the crops listed, it can commonly occur in many others including perennial fruit (especially citrus), annual field and vegetable crops, ornamentals, grassland and forestry.

Biology and Ecology

Top of page
S. viridis is an annual plant, reproducing only by seed. Freshly shed seed may be capable of germinating immediately (Holm et al., 1977), or there may be a short period of dormancy of some weeks or months, readily broken by moist storage for a few weeks (Douglas et al., 1985). Optimum temperatures for germination are 20-35°C. Germination is very much slower at 15°C and almost completely prevented at 10°C. Light is not necessary, but it has been observed that germination in natural light is higher than in light filtered through a plant canopy. Most germination occurs from the top 1-2 cm soil layer, but it can occur down to 8 or even 10 cm depth. The coleoptile is only 1 cm long, but extension of the mesocotyl (the internode below the coleoptile) allows normal emergence from greater depths. Seeds can retain their viability in soil for as long as 15-21 years, with longevity increasing with depth of burial (Douglas et al., 1985).

Emergence in the USA occurs mainly in April and May, while in Canada it is predominantly in late May, although it can continue throughout the summer.

Development of S. viridis seedlings is critically affected by light and temperature. As a C4 plant, S. viridis benefits from high temperatures and full sunlight, and is sensitive to shading, which greatly reduces tillering and seed production (Douglas et al., 1985). It also has the potential to benefit from increasing levels of carbon dioxide (Ziska and Bunce, 1997). There have been various studies of the growth of S. viridis under different conditions of light, temperature, moisture and nutrient level (see Douglas et al., 1985) and a growth model has been developed for the var. robusta-purpurea in the US mid-west (Schroll and Schreiber, 1985).

S. viridis is not profoundly affected by daylength, but does behave as a quantitative short-day plant, such that, at 22.5°C, flowering occurs after 26 days growth in an 8-hour photoperiod and after 62 days in a 16-hour photoperiod. The differences are smaller at a higher temperature of 30°C. As growth is considerably more vigorous under longer days, the weed is able to tiller and produce abundant seed within 2-3 months under the relatively long days of the temperate summer of North America (Douglas et al., 1985).

Mycorrhizal associations are believed to be important in the early stages of growth (Douglas et al., 1985).

Cultural practices have some influence on the abundance of S. viridis, with a tendency for reduced tillage to increase populations of the weed, it can, however, persist and be troublesome in most systems. In some areas S. viridis is associated with light and coarse-textured soils, but in others, it occurs on all soils including black clays. It is favoured by high nitrogen levels (Douglas et al., 1985).

There is no specialized mechanism for seed dispersal, but long-distance spread is known to have occurred through contaminated crop seed. Seeds are able to survive passage through the digestive systems of livestock and transmission with irrigation or flood water (Douglas et al., 1985; Holm et al., 1977).

Natural enemies

Top of page
Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Alternaria alternata Pathogen
Alternaria japonica Pathogen
Cosmopterix setariella
Lema concinnipennis
Magnaporthe oryzae Pathogen
Meromyza saltatrix
Oligonychus indicus
Pseudomonas fluorescens Antagonist
Pseudomonas syringae Pathogen
Pythium debaryanum Pathogen
Pythium graminicola Pathogen

Notes on Natural Enemies

Top of page
Douglas et al. (1985) provide a long list of insects associated with S. viridis (detected by sweeping or aspiration) but it is not clear how many of these are feeding on the weed. None has been seriously considered as a biological control agent.


Top of page
Holm et al. (1979) list S. viridis as a serious or principal weed in seven countries, including the USA, Canada and Japan, where it occurs in a wide range of annual and perennial crops (see Host Range). Heap and Morrison (1996) comment that S. viridis is one of the two (with Avena fatua) most abundant grass weeds of crop land in the Canadian prairie provinces. Holm et al. (1977) observe that 'its importance as a weed relates to its heavy seed production and dense competitive stands which occur largely in spring-sown crops'. Individually, S. viridis plants may not be highly competitive, but population densities can reach 3000 plants/m² (Douglas et al., 1985). The problem caused by S. viridis is also now aggravated by the evolution of herbicide-resistant biotypes. One population already shows dual resistance to both ACC-ase inhibitors and the dinitroaniline herbicides. There are no completely effective alternative herbicides available to control S. viridis.

The competitive and yield reducing effects of S. viridis depend on the associated crop, the weed density, the time of emergence, and environmental conditions (Douglas et al., 1985). Yield reductions in cereals in Canada vary greatly from season to season and depend especially on temperature early in the crop season. When wheat was planted in early May in Saskatchewan, Canada, even 1550 S. viridis plants/m² failed to affect wheat yields (Rahman and Ashford, 1972), however, in other circumstances, 100 plants/m² can reduce yields (Blackshaw et al., 1981). In the USA, wheat yield losses ranged from 0-47% when infested with 720 plants/m² (Peterson and Nalewaja, 1991). S. viridis is most competitive when it emerges with or shortly after the wheat crop (Blackshaw et al., 1981; O'Donovan, 1994). Peterson and Nalewaja (1992) showed that at 30°C, S. viridis sown 4 days before and 4 days after wheat reduced crop growth by 50 and 13%, respectively. S. viridis can be especially damaging when sowing of cereals is deliberately delayed as a means of reducing infestations of Avena fatua. In these circumstances, S. viridis is more likely to experience the high temperature conditions under which it can develop rapidly and outgrow the crop.

Models have been developed to help understand and predict competitive effects in wheat (for example, see Maxwell, 1992) and in other crops (McGiffen et al., 1997). The latter authors note that both maize and soyabean can suffer heavy losses due to competition from the 'robust' forms of S. viridis.

In maize, densities of 20 and 56 plants/m² failed to reduce yields in two trials but in the same trials densities above 40 and 89 plants/m² reduced yields by 6-18%. The competitive effects of S. viridis and S. pumila in maize were reduced with nitrogen application (Douglas et al., 1985).

In soyabean, no significant yield loss was detected from populations up to 800 plants/m² in either of two years in Italy (Sartorato et al., 1996) whereas in the USA, S. viridis var. rubusta-purpurea reduced yields by 30-63% at densities of 70-170 plants/m² (Schroll and Schreiber, 1983).

In sugarbeet, 26 and 52 plants/m², reduced yields by 27 and 36%, respectively (Douglas et al., 1985). Mesbah et al. (1994) determined the thresholds for reduction in yields of sugarbeet to be 0.06 plants of S. viridis per m of row all-season, or three plants per m of row for 3.5 weeks after crop emergence.

Douglas et al. (1985) provide further examples of the competitive effects of S. viridis, but do not comment on the likelihood of considerable differences in competitiveness between the different varieties of the species. It seems most probable that the 'giant' and 'robust' forms are significantly more damaging.

Holm et al. (1977) record that there have been reports of allelopathic effects of S. viridis on cabbage seedlings.

Contamination of crop seed (leading to 'dockage') can also be a source of financial loss. In a study in Manitoba, Canada, the average number of seeds per kg of grain varied from over 4000 in rapeseed to over 10,000 in barley (Douglas et al., 1985).


Top of page
Holm et al. (1977) indicate that S. viridis 'is sometimes used for pasture' but the extent and importance of this use is uncertain. Douglas et al. (1985) record that the seeds have approximately the same nutritive value as cereal grains. Together with S. pumila, they may make up 50% or more of the diet of some wild birds in the USA.

An interesting by-product from the development of triazine-resistance in S. viridis has been the deliberate transfer of this resistance into the crop S. italica (Italian millet) in France, so that the crop can then be safely treated with triazine herbicides (Naciri et al., 1992). It has, however, been shown that natural outcrossing can occur with S. viridis, so, even where triazine-resistance does not already occur in the weed, it is likely to develop by outcrossing from the crop (Darmency et al., 1992). Resistances to trifluralin and to sethoxydim have also been transferred from S. viridis to S. italica (Wang et al., 1997a, b).

Uses List

Top of page

Animal feed, fodder, forage

  • Forage

Human food and beverage

  • Cereal

Similarities to Other Species/Conditions

Top of page
S. italica (the crop Italian millet) is closely related to, and thought to have been derived from S. viridis. It differs in having a larger, more lobed, inflorescence, up to 3 cm wide and spikelets which break up below the upper lemma, leaving lower lemma and glumes attached. The seed is smooth, not ridged.

S. verticillata is also closely related but is normally clearly distinguished by the retrorse (downward pointing) barbs on the bristles, hence its 'sticky' inflorescence. The inflorescence is also more lobed, with spikelets grouped into whorls. This character helps to distinguish the non-sticky S. verticillata var. ambigua from S. viridis.

S. pumila is superficially very similar in form to S. viridis but differs in having more bristles per spikelet (at least five) which are yellow or reddish, not green. The upper glume is much shorter than the upper lemma, exposing the coarsely rugose upper lemma. Holm et al. (1997) provide a useful drawing comparing S. viridis with S. glauca [S. pumila] and S. faberi.

S. parviflora is close to S. pumila but differs in being perennial with a distinct rhizome.

S. faberi is more robust than typical S. viridis, and has larger spikelets, 2.5 to 3 mm long, more bristles (2-6 per spikelet) and a larger, nodding inflorescence up to 2 cm wide. It is often confused with S. viridis var. major which also has a nodding inflorescence, but Parochetti (1973) points out that S. faberi has an abundance of short hairs on the upper leaf surface, while S. viridis var. major is only rough to the touch. Also the seed heads of S. faberi are whitish-yellow at maturity while those of S. viridis var. major are reddish-purple. Schreiber and Oliver (1971) provide a useful key and other details for distinguishing the various forms of S. viridis from S. faberi and S. pumila as well as from each other.

Prevention and Control

Top of page

Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.

Cultural Control

S. viridis, being a tufted annual grass, is readily controlled by all normal tillage practices. Where these do not suffice and other non-chemical methods are needed, early planting is one of the main recommended practices for reducing S. viridis importance in cereals in North America, as wheat is able to establish at lower temperatures than the weed (Douglas et al., 1985; Khan et al., 1996). Dense planting and increased nutrient are also helpful in wheat and barley. Higher density is also suggested for growing maize without herbicide in Ukraine (Bobro et al., 1994). Crop rotation is recommended as a means of reducing the S. viridis seed bank in the USA (Jordan et al., 1995).

Chemical Control

S. viridis is normally susceptible to a very wide range of standard herbicides recommended for annual grass control. These include atrazine and other triazines, trifluralin and other dinitroanilines, metolachlor, diclofop, fluazifop, sethoxydim and other inhibitors of acetyl coenzyme A carboxylase (ACCase), propanil, EPTC and butylate, paraquat, glyphosate, glufosinate. Douglas et al. (1985) give a number of examples of herbicide use in cereals and some other crops. Williams and Schreiber (1976) comment that the giant and robust forms of S. viridis are more resistant to certain herbicides.

Herbicide Resistance

Herbicide resistance has developed quite widely, and atrazine-resistant populations are now common in Europe and the USA ( de Prado et al., 1993; Wang and Dekker, 1995). Atrazine-resistance also involves some degree of cross-resistance to related herbicides. Resistance to the ACCase inhibitors, and to trifluralin has also developed in North America. Resistance to trifluralin has been associated with cross-resistance to all other dinitroaniline herbicides, and to some other herbicides inhibiting mitosis including chlorthal dimethyl and dithiopyr (Beckie and Morrison, 1993; McAlister et al., 1995). This resistance was thought to be controlled by a single recessive gene (Jasieniuk et al., 1994) but is now believed to involve a complex of genes (Wang et al., 1996). Resistance to trifluralin is not associated with any reduction in fitness, and resistant populations can persist for at least 7 years even in the absence of selection pressure for herbicide resistance (Andrews and Morrison, 1997). Dual resistance to trifluralin and ACCase inhibitors has now been detected in Canada (Heap and Morrison, 1996). In the case of ACCase inhibitors, the cross-resistance pattern is complex, suggesting that there have been a number of different mutations affecting the sensitivity of the enzyme in different populations (Shukla et al., 1997). Resistance to sethoxydim is apparently controlled by a single dominant gene (Wang et al., 1997a).


Top of page

Andrews TS, Morrison IN, 1997. The persistence of trifluralin resistance in green foxtail (Setaria viridis) populations. Weed Technology, 11(2):369-372; 12 ref.

Auld BA, Medd RW, 1987. Weeds. An illustrated botanical guide to the weeds of Australia. Melbourne, Australia; Inkata Press, 255 pp.

Beckie HJ, Morrison IN, 1993. Effect of ethalfluralin and other herbicides on trifluralin-resistant green foxtail (Setaria viridis). Weed Technology, 7(1):6-14; 24 ref.

Blackshaw RE, Stobbe EH, Shaykewich CF, Woodbury W, 1981. Influence of soil temperature and soil moisture on green foxtail (Setaria viridis) establishment in wheat (Triticum aestivum). Weed Science, 29(2):179-184

Bobro MA, Bachassi A, 1994. Sowing date, plant density and hybrid as the basis of technology for growing maize without herbicides. Selektsionno-geneticheskie i biotekhnologicheskie priemy povysheniya produktivnosti sel'skokhozyaistvennykh rastenii., 77-82; 8 ref.

Chaudhary SA, Parker C, Kasasian L, 1981. Weeds of Central, Southern and Eastern Arabian Peninsula. Tropical Pest Management, 27(2):181-190.

Clayton WD, 1972. Gramineae. In: Hutchinson J, Dalziel JM, Hepper FN, 1972. Flora of West Tropical Africa. Vol 3. Part 2. London, UK: Crown Agents, 349-512.

Clayton WD, 1980. Setaria. In: Tutin TG, Heywood VH, Burges NA, Moore DM, Valentine DH, Walters SM, Webb DA, eds. Flora Europaea, Volume 5. Alismataceae to Orchidaceae Monocotyledones. Cambridge, UK: Cambridge University Press, 263-264.

Clayton WD, Renvoize SA, 1982. Flora of Tropical East Africa. Graminea (Part 3). Rotterdam, The Netherlands: A.A. Balkema, 448 pp.

Darmency H, Lefol E, Chadoeuf R, 1992. Risk assessment of the release of herbicide resistant transgenic crops: two plant models. IXe Colloque international sur la biologie des mauvaises herbes, 16-18 September 1992, Dijon, France., 513-523; 25 ref.

Dmitrieva SA, 1985. Chromosome numbers of representatives of the families Lamiaceae and Poaceae in the Byelorussian flora. Botanicheskii Zhurnal, 70:128-130.

Douglas BJ, Thomas AG, Morrison IN, Maw MG, 1985. The biology of Canadian weeds. 70. Setaria viridis (L.) Beauv. Canadian Journal of Plant Science, 65(3):669-690; [2 fig.]; 6 pp. of ref.

Erik S, Demirkus, 1988. New localities for some plants in the flora of Turkey. Doga, Turk Botanik Dergisi, 12:224-233.

Flora do Brasil 2020, 2017. Website under construction. Brazil: Jardim Botânico do Rio de Janeiro.

Hafliger E, Scholz H, 1980. Grass weeds I. Weeds of the subfamily Panicoideae. Basle, Switzerland: Ciba-Geigy Ltd.

Heap IM, Morrison IN, 1996. Resistance to aryloxyphenoxypropionate and cyclohexanedione herbicides in green foxtail (Setaria viridis). Weed Science, 44(1):25-30; 15 ref.

Holm LG, Doll J, Holm E, Pancho JV, Herberger JP, 1997. World Weeds: Natural Histories and Distribution. New York, USA: John Wiley & Sons Inc.

Holm LG, Pancho JV, Herberger JP, Plucknett DL, 1979. A geographical atlas of world weeds. New York, USA: John Wiley and Sons, 391 pp.

Holm LG, Plucknett DL, Pancho JV, Herberger JP, 1977. The World's Worst Weeds. Distribution and Biology. Honolulu, Hawaii, USA: University Press of Hawaii.

Jasieniuk M, Brûlé-Babel AL, Morrison IN, 1994. Inheritance of trifluralin resistance in green foxtail (Setaria viridis). Weed Science, 42(1):123-127; 27 ref.

Jordan N, Mortensen DA, Prenzlow DM, Cox KC, 1995. Simulation analysis of crop rotation effects on weed seedbanks. American Journal of Botany, 82(3):390-398; 24 ref.

Kaul MK, 1986. Weed Flora of Kashmir Valley. Jodhpur, India: Scientific Publishers, 422 pp.

Keresztes Z, Dorner Z, Zalai M, 2014. Weed composition and diversity of three organic farms in Hungary. IOBC/WPRS Bulletin [Proceedings of the IOBC/WPRS Working Group "Landscape Management for Functional Biodiversity", Poznan, Poland, 21-31 May 2014.], 100:69-72.

Khan M, Donald WW, Prato T, 1996. Spring wheat (Triticum aestivum) management can substitute for diclofop for foxtail (Setaria spp.) control. Weed Science, 44(2):362-372; 23 ref.

Litvinov PI, Grushkova SA, Syrel'shchikova DP, Chebanovskaya AF, 1987. Chemical control in grapevine rootstock nurseries. Zashchita Rastenii, No.3:32.

Lorenzi HJ, Jeffery LS(Editors), 1987. Weeds of the United States and their control. New York, USA; Van Nostrand Reinhold Co. Ltd., 355 pp.

Maxwell BD, 1992. Weed thresholds: the space component and considerations for herbicide resistance. Weed Technology, 6(1):205-212; [presented at a symposium on the ecological perspectives on utility of thresholds for weed management held in Louisville, USA, 5 February 1991]; 30 ref.

McAlister FM, Holtum JAM, Powles SB, 1995. Dinitroaniline herbicide resistance in rigid ryegrass (Lolium rigidum). Weed Science, 43(1):55-62; 22 ref.

McGiffen MEJr, Forcella F, Lindstrom MJ, Reicosky DC, 1997. Covariance of cropping systems and foxtail density as predictors of weed interference. Weed Science, 45(3):388-396; 30 ref.

Mesbah A, Miller SD, Fornstrom KJ, Legg DE, 1994. Kochia (Kochia scoparia) and green foxtail (Setaria viridis) interference in sugarbeets (Beta vulgaris). Weed Technology, 8(4):754-759; 10 ref.

Mokshin VS, 1986. Effectiveness of herbicides depending on moisture conditions and methods of soil cultivation. Sibirskii Vestnik Sel'skokhozyaistvennoi Nauki, No.5:8-12; 3 ref.

Naciri Y, Darmency H, Belliard J, Dessaint F, Pernès J, 1992. Breeding strategy in foxtail millet, Setaria italica (L. P. Beauv.) following interspecific hybridization. Euphytica, 60(2):97-104; 10 ref.

Numata M, Yoshizawa N, 1975. Weed flora of Japan. Japan Association for the Advancement of Phyto-Regulators. Tokyo, Japan: Zenkoku Noson Kyoiku Kyokai.

O'Donovan JT, 1994. Green foxtail (Setaria viridis) and pale smartweed (Polygonum lapathifolium) interference in field crops. Weed Technology, 8(2):311-316; 16 ref.

Parochetti JV, 1973. Giant green foxtail in Maryland and surrounding areas. Proceedings of the Northeastern Weed Science Society, New York, 1973. Volume 27, 168-169.

Peterson DE, Nalewaja JD, 1991. Green foxtail (Setaria viridis) competition with spring wheat (Triticum aestivum). Weed Technology, 6(2):291-296; 9 ref.

Peterson DE, Nalewaja JD, 1992. Environment influences green foxtail (Setaria viridis) competition with wheat (Triticum aestivum). Weed Technology, 6(3):607-610; 14 ref.

Prado Rde, Romero E, Tena M, 1993. Chloroplastic susceptibility of three Setaria species to different photosynthesis-inhibiting herbicides. Proceedings of the 1993 Congress of the Spanish Weed Science Society, Lugo, Spain, 1-3 December 1993., 239-242; 5 ref.

Rahman A, Ashford R, 1972. Control of green foxtail in wheat with trifluralin. Weed Science, 20(1):23-27

Reflora - Virtual Herbarium, 2017. Brazil: Jardim Botânico do Rio de Janeiro.

Sartorato I, Berti A, Zanin G, 1996. Estimation of economic thresholds for weed control in soybean (Glycine max (L.) Merr.). Crop Protection, 15:63-68.

Schreiber MM, Oliver LR, 1971. Two new varieties of Setaria viridis. Weed Science, 19:424-427.

Schroll RE, Schreiber MM, 1983. Growth analysis of robust purple foxtail interference in soybean. Proceedings, North Central Weed Control Conference., 34-35.

Schroll RE, Schreiber MM, 1985. Sensitivity analysis of SETSIM (Setaria simulation). Proceedings, North Central Weed Control Conference., Vol.40:6-7.

Shukla A, Leach GE, Devine MD, 1997. High-level resistance to sethoxydim conferred by an alteration in the target enzyme, acetyl-CoA carboxylase, in Setaria faberi and Setaria viridis. Plant Physiology and Biochemistry (Paris), 35(10):803-807; 20 ref.

Shukla U, 1996. The Grasses of North-Eastern India. Jodhpur, India: Scientific Publishers, 325 pp.

Species Link, 2017. Multiple data sets. Brazil: Center for Reference in Environmental Information.

Stace C, 1991. New Flora of the British Isles. Cambridge, UK: Cambridge University Press.

Teerawatsakul M, Takayanagi S, Kusanagi T, Noda K, 1987. Characteristics of seed germination of Euphorbia geniculata, an upland weed in Thailand. Weed Research, Japan, 32(3):168-172; 6 ref.

Tselev NN, Fedorov AA, 1983. Grasses of the Soviet Union. Part 2. New Delhi, India: Oxonian Press Ltd.

Wang RL, Dekker J, 1995. Weedy adaptation in Setaria spp. III. Variation in herbicide resistance in Setaria spp. Pesticide Biochemistry and Physiology, 51(2):99-116; 52 ref.

Wang RL, Wendel JF, Dekker JH, 1995. Weedy adaptation in Setaria spp. I. Isozyme analysis of genetic diversity and population genetic structure in Setaria viridis. American Journal of Botany, 82(3):308-317; 44 ref.

Wang T, Darmency H, Wang TY, 1997. Dinitroaniline herbicide cross-resistance in resistant Setaria italica lines selected from interspecific cross with S. viridis. Pesticide Science, 49:277-283.

Wang T, Fleury A, Ma J, Darmency H, 1996. Genetic control of dinitroaniline resistance in foxtail millet (Setaria italica). Journal of Heredity, 87(6):423-426; 17 ref.

Wang TianYu, Darmency H, 1997. Inheritance of sethoxydim resistance in foxtail millet, Setaria italica (L.) Beauv. Euphytica, 94(1):69-73; 19 ref.

Wang ZM, Devos KM, Liu CJ, Wang RQ, Gale MD, 1998. Construction of RFLP-based maps of foxtail millet, Setaria italica (L.) P. Beauv. Theoretical and Applied Genetics, 96:31-36.

Wang ZR, 1990. Farmland Weeds in China. Beijing, China: Agricultural Publishing House.

William RD, Schreiber MM, 1976. Numerical and chemotaxonomy of the green foxtail complex. Weed Science, 24(3):331-335

Ziska LH, Bunce JA, 1997. Influence of increasing carbon dioxide concentration on the photosynthetic and growth stimulation of selected C crops and weeds. Photosynthesis Research, 54(3):199-208; 25 ref.

Distribution References

Anon, 1975. Weed flora of Japan (illustrated by colour). In: Weed flora of Japan (illustrated by colour). [ed. by Numata M, Yoshizawa N]. Tokyo, Japan: Japan Association for the Advancement of Phyto-Regulators. 415 pp.

Anon, 1987. Weeds of the United States and their control. [ed. by Lorenzi H J, Jeffery L S]. New York, USA: Van Nostrand Reinhold Co. Ltd. 355 pp.

Anon, 1987. Weeds. An illustrated botanical guide to the weeds of Australia. [ed. by Auld B A, Medd R W]. Melbourne, Australia: Inkata Press. 255 pp.

CABI, Undated. Compendium record. Wallingford, UK: CABI

Celepcİ E, Uygur S, Kaydan M B, Uygur F N, 2017. Mealybug (Hemiptera: Pseudococcidae) species on weeds in Citrus (Rutaceae) plantations in Çukurova Plain, Turkey. Türkiye Entomoloji Bülteni. 7 (1), 15-21.

Chaudhary S A, Parker C, Kasasian L, 1981. Weeds of central, southern and eastern Arabian Peninsula. Tropical Pest Management. 27 (2), 181-190.

Clayton WD, 1972. Gramineae. In: Flora of West Tropical Africa, 3 (2) [ed. by Hutchinson J, Dalziel JM, Hepper FN]. London, UK: Crown Agents. 349-512.

Clayton WD, 1980. Setaria. In: Flora Europaea. Alismataceae to Orchidaceae Monocotyledones, 5 [ed. by Tutin TG, Heywood VH, Burges NA, Moore DM, Valentine DH, Walters SM, Webb DA]. Cambridge, UK: Cambridge University Press. 263-264.

Dmitrieva S A, 1985. Chromosome numbers of representatives of the families Lamiaceae and Poaceae in the Byelorussian flora. Botanicheskiĭ Zhurnal. 70 (1), 128-130.

Douglas B J, Thomas A G, Morrison I N, Maw M G, 1985. The biology of Canadian weeds. 70. Setaria viridis (L.) Beauv. Canadian Journal of Plant Science. 65 (3), 669-690.

Evans C K, Bag S, Frank E, Reeve J, Ransom C, Drost D, Pappu H R, 2009. Green foxtail (Setaria viridis), a naturally infected grass host of Iris yellow spot virus in Utah. Plant Disease. 93 (6), 670-671. DOI:10.1094/PDIS-93-6-0670C

Fazal Hadi, Muhammad Ibrar, 2015. Ecology of weeds in wheat crops of Kalash valley, district Chitral, Hindukush Range, Pakistan. Pakistan Journal of Weed Science Research. 21 (3), 425-433.

Golnaraghi A R, Pourrahim R, Farzadfar S, Ohshima K, Shahraeen N, Ahoonmanesh A, 2007. Incidence and distribution of Tomato yellow fruit ring virus on soybean in Iran. Plant Pathology Journal (Faisalabad). 6 (1), 14-21.

Hassannejad S, Ghafarbi S P, 2013. Weed flora survey of Tabriz wheat (Triticum aestivum L.) fields. Journal of Biodiversity and Environmental Sciences (JBES). 3 (9), 118-132.

Hassannejad S, Ghafarbi S P, Abbasvand E, Ghisvandi B, 2014. Quantifying the effects of altitude and soil texture on weed species distribution in wheat fields of Tabriz, Iran. Journal of Biodiversity and Environmental Sciences (JBES). 5 (1), 590-596.

Hassannejad S, Ghisvandi B, 2013. Grasses distribution in wheat fields of Tabriz-Iran and recorded Sclerochloa woronowii (Hack.) Tzvelev as a new weed species for flora of Iran. Technical Journal of Engineering and Applied Sciences. 3 (22), 3119-3124.

He YunHe, Qiang Sheng, 2014. Analysis of farmland weeds species diversity and its changes in the different cropping systems. Bulgarian Journal of Agricultural Science. 20 (4), 786-794.

Holm L, Pancho J V, Herberger J P, Plucknett D L, 1979. A geographical atlas of world weeds. New York, Chichester (), Brisbane, Toronto, UK: John Wiley and Sons. xlix + 391 pp.

Hwang KiSeon, Eom MinYong, Park SuHyuk, Won OkJae, Lee InYong, Park KeeWoong, 2015. Occurrence and distribution of weed species on horticulture fields in Chungnam province of Korea. Journal of Ecology and Environment. 38 (3), 353-360. DOI:10.5141/ecoenv.2015.036

Ibrahim I K A, Mokbel A A, Handoo Z A, 2010. Current status of phytoparasitic nematodes and their host plants in Egypt. Nematropica. 40 (2), 239-262.

Kaul M K, 1986. Weed flora of Kashmir Valley. Jodhpur, India: Scientific Publishers. 422pp.

Keresztes Z, Dorner Z, Zalai M, 2014. Weed composition and diversity of three organic farms in Hungary. IOBC/WPRS Bulletin. 69-72.

Litvinov P I, Grushkova S A, Syrel'shchikova D P, Chebanovskaya A F, 1987. Chemical control in grapevine rootstock nurseries. Zashchita Rasteniĭ. 32.

Mokshin V S, 1986. Effectiveness of herbicides depending on moisture conditions and methods of soil cultivation. Sibirskiĭ Vestnik Sel'skokhozyaĭstvennoĭ Nauki. 8-12.

Moskova T, Dimitrov G, Tityanov M, 2018. Distribution and degree of weed growth of amaranth and other weeds in sunflower crops in Plovdiv and Stara Zagora regions. Journal of Mountain Agriculture on the Balkans. 21 (1), 158-168.

Reflora - Virtual Herbarium, 2017. (Jardim Botânico do Rio de Janeiro)., Brazil:

Shukla U, 1996. The Grasses of North-Eastern India., Jodhpur, India: Scientific Publishers. 325 pp.

Species Link, 2017. Multiple data sets., Brazil: Center for Reference in Environmental Information.

Teerawatsakul M, Takayanagi S, Kusanagi T, Noda K, 1987. Characteristics of seed germination of Euphorbia geniculata, an upland weed in Thailand. Weed Research, Japan. 32 (3), 168-172.

Vafaee B S, Narimani V, Farokhzad A, Chasemzadeh R, 2011. Quantitative evaluation of predominant of weeds in winter wheat and barley fields in Eastern Azerbaijan, Iran. Revista Cientifica UDO Agricola. 11 (1), 126-133.

Vojnich V J, Pölös E, Baglyas F, 2017. Weed vegetation of a vineyard on sandy soil. Lucrări Științifice, Universitatea de Științe Agricole Și Medicină Veterinară a Banatului, Timisoara, Seria I, Management Agricol. 19 (1), 119-122.

Vrbničanin S, Božić D, Sarić M, Pavlović D, Matić L, Dakić P, 2012. Biological spectrum of weed flora and vegetation of raspberry plantings in Serbia. Acta Horticulturae. 293-296.

Distribution Maps

Top of page
You can pan and zoom the map
Save map
Select a dataset
Map Legends
  • CABI Summary Records
Map Filters
Third party data sources: