Eichhornia crassipes (water hyacinth)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Plant Type
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Host Plants and Other Plants Affected
- Biology and Ecology
- Air Temperature
- Soil Tolerances
- Water Tolerances
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Impact Summary
- Economic Impact
- Environmental Impact
- Threatened Species
- Social Impact
- Risk and Impact Factors
- Uses List
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Links to Websites
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Eichhornia crassipes (Mart.) Solms 1883
Preferred Common Name
- water hyacinth
Other Scientific Names
- Eichhornia cordifolia Gandoger 1920
- Eichhornia crassicaulis Schlect. 1862
- Eichhornia speciosa Kunth 1843
- Heteranthera formosa Miq. 1843
- Piaropus crassipes (Mart.) Raf. 1837
- Piaropus mesomelas Raf. 1837
- Pontederia crassicaulis Schlect. 1862
- Pontederia crassipes Mart. 1823
- Pontederia elongata Balf. 1855
International Common Names
- English: floating water hyacinth; lilac devil; Nile lily; pickerelweed; water orchid; water violet
- Spanish: aguapey (Argentina); cola de pato; hierba jicotea; lagunero (Nicaragua); lechuguilla; lila de agua; lila de caño; pontederia azul (Mexico); reina del agua; taruya (Nicaragua)
- French: bofinace; héliotrope; jacinthe d'eau
- Portuguese: jacinto aquatico
Local Common Names
- Antigua and Barbuda: water violet
- Argentina: aquapey; camalotes; jacinto de agua
- Bangladesh: kachuripana
- Brazil: aguape de flor roxa; aguape puru-a; baronesa; dama del lago; jacinta d'agua; murumurii
- Cambodia: kamplauk
- Chile: jiro de agua; violeta de agua
- Colombia: buchon; lirio de agua; tarulla
- Congo: kongo ya sika
- Costa Rica: lirio de agua
- Cuba: boniatillo de agua; flor de agua; hierba de jicotea; jacinto de agua; lirio acauático; malangueta
- Czechoslovakia (former): tokozelka; vodin hyacint
- Denmark: vanhyazint
- Dominican Republic: lila de agua
- Egypt: bisnidh; habba; halassandi/halassant; war-el-nil; zanim; zoqqeym et-tani
- El Salvador: halsa; lechugo; lechugo de concha
- Fiji: babadabeniga; bekabekairaga; jalkhumbe
- Former USSR: wampee
- France: eichhornie
- Germany: wasserhyazinthe
- Guatemala: lirio acuatico; ninfa
- India: akasa thamarai; German pana; jalkhumbi; kachuripana; kajor pati; kolavazha; kulavali; neithamarai; pisachi thanana; sokh-samundar; tagoi; vilayati pana
- Indonesia: bengai gondo; bengok; bia bia; eceng; eceng gondok; eceng padi; gendot; ilung ilung; mampau/mampoh; nappong; sekar bopong; wewehan
- Israel: yakinton hamaim
- Italy: giacinto d'acqua
- Jamaica: water lily
- Japan: hotei-aoi; torin; uchikusa; weinchan
- Lesser Antilles: glaïeul bleu
- Madagascar: tetezanalika; tsikafokafona
- Malaysia: bunga jamban; keladi bunting; kemeling telur
- Mauritius: hoteiaoi
- Mexico: jacinto acuatico; lirio acuatico
- Myanmar: beda-bin; ye-padauk
- Netherlands: waterhyacint
- Nicaragua: lirio de agua
- Pakistan: gulbakauli; kalali
- Peru: camalote; lirio de agua
- Philippines: water lily
- Puerto Rico: flor de agua
- South Africa: Florida devil; lilac devil; waterhiasint
- Spain: lirio de agua
- Sri Lanka: diya kehel; diya manel; habara/habarala; sabara; yapura
- Suriname: badawaro; moessiri; oponopa-joelire
- Taiwan: putailien
- Thailand: paktopjava; sawah; top-chawa
- Turkey: su sümbülü
- Uruguay: aguape/aguape-puru
- USA: river raft
- Venezuela: bora; lagunera
- Vietnam: luc-binh
- EICCR (Eichhornia crassipes)
Summary of InvasivenessTop of page
E. crassipes, a native of South America, is a major freshwater weed in most of the frost-free regions of the world and is generally regarded as the most troublesome aquatic plant (Holm et al., 1997). It has been widely planted as a water ornamental around the world because of its striking flowers. Wherever it has encountered suitable environmental conditions it has spread with phenomenal rapidity to form vast monotypic stands in lakes, rivers and rice paddy fields. Then it adversely affects human activities (fishing, water transport) and biodiversity. It is impossible to eradicate, and often only an integrated management strategy, inclusive of biological control, can provide a long-term solution to this pest.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Plantae
- Phylum: Spermatophyta
- Subphylum: Angiospermae
- Class: Monocotyledonae
- Order: Pontederiales
- Family: Pontederiaceae
- Genus: Eichhornia
- Species: Eichhornia crassipes
Notes on Taxonomy and NomenclatureTop of page
Although almost certainly collected as early as 1801 in Colombia, the species was first described in 1824 and given the name Pontederia crassipes by C.F.P. von Martius from specimens collected in Brazil. Kunth in 1843 split the genus and created Eichhornia to cover species with trilocular ovary and numerous ovules. He ignored the epithet 'crassipes' and used the name Eichhornia speciosa Kunth. He also ignored Rafinesque's revision of 1836 in which the genus had been given the name Piaropus. A number of other combinations were applied by different authors in the nineteenth century, but finally, in 1883, H. Solms-Laubach established the combination Eichhornia crassipes (Mart.) Solms by which the species is now universally known. This ignores the priority of the name Piaropus on the basis that Eichhornia had been in use since 1843 and was regarded as a nomen conservatum (after Gopal, 1987).
In early years there was considerable confusion with the closely related Eichhornia azurea (Swartz) Kunth, which had been collected and described as Pontederia azurea Sw. somewhat earlier in 1797. The distribution of this species overlaps with that of Eichhornia crassipes in South and Central America. Eichhornia azurea differs in having finely toothed petals, a more elongated main stem (not spreading by stolons) and distichous leaves lacking swollen petioles. Even now there is confusion between the two species in some areas, resulting from excessive reliance on the petiole character and an assumption that a lack of swollen petioles means it must be Eichhornia azurea.
Six other species of Eichhornia have been described, mainly from South and Central America but including Eichhornia natans (P. Beauv.) Solms. which is restricted to Africa. All are relatively rare and of little or no economic importance. Confusion with Eichhornia natans is unlikely as leaves of the latter rarely exceed 4 cm long and flowers are less than 2 cm across.
DescriptionTop of page
The initial leaves of seedling E. crassipes are elongated and strap-like, but soon develop the familiar spathulate form and, under suitable unshaded conditions, swollen petioles which ensure that, once dislodged, the seedlings will float from the mud into open water. The plant is very variable in size, seedlings having leaves that are only a few centimetres across or high, whereas mature plants with good nutrient supply may reach 1 m in height. Plants in an uncrowded situation tend to have short, spreading petioles with pronounced swelling, while in a dense stand they are taller, more erect and with little or no swelling of the petioles.
The plant system consists of individual shoots/crowns each with up to ten expanded leaves arranged spirally (3/8 phyllotaxy) and separated by very short internodes. As individual shoots develop, the older leaves die off leaving a stub of leafless dead shoot projecting downwards. This may eventually cause the whole shoot to sink and die.
Leaves consist of petiole (often swollen, 2-5 cm thick) and blade (roughly round, ovoid or kidney-shaped, up to 15 cm across). The base of the petiole and any subsequent leaf is enclosed in a stipule up to 6 cm long.
Roots develop at the base of each leaf and form a dense mass: usually 20-60 cm long, though they can extend to 300 cm. The ratio of root to shoot depends on the nutrient conditions, and in low nutrient conditions they may account for over 60% of the total plant weight. They are white when formed in total darkness but often purplish under field conditions, especially in conditions of low nutrients.
Periodically, axillary buds develop as stolons, growing horizontally for 10-50 cm before establishing daughter plants. Extremely large populations of inter-connected shoots can develop very rapidly, though the connecting stolons eventually die.
The inflorescence is a spike which develops from the apical meristem, but tends to appear lateral owing to the immediate development of an axillary bud as a 'renewal' or 'continuation' shoot. Each spike, up to 50 cm high, is subtended at the base by two bracts and has 8-15 sessile flowers (rarely 4-35). Each flower has a perianth tube 1.5 cm long, expanding into six mauve or purple lobes up to 4 cm long. The main lobe has a bright-yellow, diamond-shaped patch surrounded by deeper purple. Once the inflorescence is fully emerged from the leaf sheath, flowers all open together, starting at night, completing the process in the morning and withering by the next night when the peduncle starts to bend down. Each capsule may contain up to 450 small seeds, each about 1 x 3 mm.
The flowers are tristylous. They have six stamens and one style, arranged in three possible configurations (floral trimorphism) - with short style (and medium and long stamens), medium style (short and long stamens) or long style (short and medium stamens). The medium style form is genetically dominant and is by far the commonest form in almost all infested areas. The short-styled form is only known from South America, whereas the long-styled form is found commonly in South America, more rarely in South-East Asia and very rarely in Africa. Only in Sri Lanka is the long-styled the commonest form. Some other tristylous species show incompatibility between the different forms but E. crassipes does not. Hence pollination (mainly by wind) can result in good seed set, though in some populations there may be a higher degree of self-incompatibility.
Plant TypeTop of page Aquatic
DistributionTop of page
E. crassipes originated in tropical South America, but is now naturalized in Africa, Australia, India and many other countries.
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Bangladesh||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Bhutan||Restricted distribution||Introduced||Parker, 1992|
|Brunei Darussalam||Present||Introduced||Waterhouse, 1993; EPPO, 2014|
|Cambodia||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|China||Widespread||Introduced||1901||Invasive||Xie et al., 2001; EPPO, 2014|
|-Fujian||Widespread||Introduced||Invasive||Ding et al., 2001; EPPO, 2014|
|-Guangdong||Widespread||Introduced||Invasive||Ding et al., 2001; EPPO, 2014|
|-Hong Kong||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Jiangsu||Present||Introduced||Dai and Zhang, 1988|
|-Yunnan||Widespread||Introduced||Invasive||Ding et al., 2001; Xie et al., 2001; EPPO, 2014|
|-Zhejiang||Widespread||Introduced||Invasive||Ding et al., 2001; EPPO, 2014|
|India||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Andhra Pradesh||Widespread||Introduced||Invasive||Gopal, 1987|
|-Arunachal Pradesh||Widespread||Introduced||Invasive||Gopal, 1987|
|-Himachal Pradesh||Widespread||Introduced||Invasive||Gopal, 1987|
|-Indian Punjab||Widespread||Introduced||Invasive||Gopal, 1987|
|-Madhya Pradesh||Widespread||Introduced||Invasive||Gopal, 1987|
|-Rajasthan||Restricted distribution||Introduced||Gopal, 1987|
|-Tamil Nadu||Widespread||Introduced||Invasive||Gopal, 1987|
|-Uttar Pradesh||Widespread||Introduced||Invasive||Gopal, 1987|
|-West Bengal||Widespread||Introduced||Invasive||Gopal, 1987|
|Indonesia||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Irian Jaya||Widespread||Introduced||Invasive||Gopal, 1987|
|Israel||Present, few occurrences||Invasive||EPPO, 2014|
|Japan||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Korea, DPR||Present||Introduced||Dostalek et al., 1989|
|Korea, Republic of||Widespread||Introduced||Invasive||Gopal, 1987|
|Laos||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Malaysia||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Peninsular Malaysia||Widespread||Introduced||Invasive||Gopal, 1987|
|Maldives||Present||Introduced||Invasive||Pallewatta et al., 2003; EPPO, 2014|
|Myanmar||Present||Introduced||Gopal, 1987; EPPO, 2014|
|Pakistan||Restricted distribution||Introduced||Gopal, 1987; Imran et al., 2013|
|Philippines||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Singapore||Widespread||Introduced||1903||Invasive||Gopal, 1987; EPPO, 2014|
|Sri Lanka||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Taiwan||Present||Introduced||Invasive||Ding et al., 2001; EPPO, 2014|
|Thailand||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Turkey||Present||Uremis et al., 2014|
|Vietnam||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Angola||Restricted distribution||Introduced||Gopal, 1987|
|Benin||Restricted distribution||Introduced||EPPO, 2014|
|Burkina Faso||Present||EPPO, 2014|
|Burundi||Present||Introduced||Invasive||Moorhouse et al., 2001; EPPO, 2014|
|Central African Republic||Restricted distribution||Introduced||Gopal, 1987|
|Congo||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Congo Democratic Republic||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Côte d'Ivoire||Restricted distribution||Introduced||Harley, 1993; EPPO, 2014|
|Egypt||Widespread||Introduced||Invasive||Fayad et al., 2001; EPPO, 2014|
|Equatorial Guinea||Present||EPPO, 2014|
|Ethiopia||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Ghana||Present||Introduced||Invasive||EPPO, 2014; deGraft-Johnson and Akpabey, 2015; Ernest, 2015||River Oti arm of Volta Lake, Tano River and Lagoon complex, Jewi Wharf, Kpong Headpond, lower Volta River, Odaw stream in Accra|
|Kenya||Widespread||Introduced||1989||Invasive||Owiti, 1990; Mailu, 2001; IPPC-Secretariat, 2005; EPPO, 2014|
|Madagascar||Widespread||Introduced||Invasive||Binggeli, 2003; EPPO, 2014|
|Malawi||Widespread||Introduced||1960s||Invasive||Harley, 1993; Phiri et al., 2001; EPPO, 2014|
|Mali||Restricted distribution||Introduced||Lomer, 1995|
|Mauritius||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Mozambique||Restricted distribution||Introduced||Gopal, 1987; EPPO, 2014|
|Niger||Restricted distribution||Introduced||Akinyemiju, 1987; Lomer, 1995|
|Nigeria||Restricted distribution||Introduced||Invasive||Akinyemiju, 1987; EPPO, 2014|
|Réunion||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Rodriguez Island||Present||Introduced||Royal Botanic Gardens Kew, 2012|
|Rwanda||Widespread||Introduced||Invasive||Harley, 1993; EPPO, 2014|
|Senegal||Restricted distribution||Introduced||Gopal, 1987; EPPO, 2014|
|Seychelles||Present||Introduced||Royal Botanic Gardens Kew, 2012|
|Sierra Leone||Present||EPPO, 2014|
|South Africa||Widespread||Introduced||Invasive||Jones, 2001; EPPO, 2014|
|Sudan||Widespread||Introduced||1957||Invasive||Gay, 1960; Gopal, 1987; EPPO, 2014|
|Tanzania||Widespread||Introduced||Invasive||Mallya et al., 2001; EPPO, 2014|
|-Zanzibar||Restricted distribution||Introduced||Gopal, 1987|
|Uganda||Widespread||Introduced||Invasive||Hill, 1999; Mailu, 2001; EPPO, 2014; Witt and Luke, 2017|
|Zambia||Widespread||Introduced||Invasive||Bennett, 1972; Hill, 1997; EPPO, 2014; Witt and Luke, 2017|
|Zimbabwe||Widespread||Introduced||Invasive||Chikwenhere, 2001; EPPO, 2014|
|Bermuda||Widespread||Introduced||Invasive||Gopal, 1987; Kairo et al., 2003; EPPO, 2014|
|Canada||Present||Present based on regional distribution.|
|-Ontario||Present||Adebayo et al., 2011|
|Mexico||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|USA||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Alabama||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Arkansas||Present||Introduced||Center et al., 2002; EPPO, 2014|
|-California||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Florida||Widespread||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Georgia||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Hawaii||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Louisiana||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Mississippi||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-New Jersey||Present||EPPO, 2014|
|-New York||Present||EPPO, 2014|
|-North Carolina||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-South Carolina||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Texas||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
Central America and Caribbean
|Antigua and Barbuda||Present||Introduced||Acevedo-Rodríguez and Strong, 2012|
|Aruba||Present||Introduced||Royal Botanic Gardens Kew, 2012|
|Bahamas||Present||Introduced||Invasive||Gopal, 1987; Kairo et al., 2003; EPPO, 2014|
|Costa Rica||Present||Gopal, 1987; EPPO, 2014|
|Cuba||Widespread||Introduced||Invasive||Gopal, 1987; Oviedo Prieto et al., 2012; EPPO, 2014|
|Dominica||Present||Introduced||Acevedo-Rodríguez and Strong, 2012|
|Dominican Republic||Widespread||Invasive||Gopal, 1987; Kairo et al., 2003; EPPO, 2014|
|El Salvador||Present||Gopal, 1987|
|Guadeloupe||Present||Introduced||Acevedo-Rodríguez and Strong, 2012|
|Guatemala||Present||Gopal, 1987; EPPO, 2014|
|Haiti||Present||Introduced||Gopal, 1987; EPPO, 2014|
|Honduras||Present||Gopal, 1987; EPPO, 2014|
|Jamaica||Widespread||Introduced||Invasive||Gopal, 1987; Kairo et al., 2003; EPPO, 2014|
|Martinique||Present||Introduced||Acevedo-Rodríguez and Strong, 2012|
|Nicaragua||Present||Gopal, 1987; EPPO, 2014|
|Panama||Widespread||Gopal, 1987; EPPO, 2014|
|Puerto Rico||Widespread||Introduced||Invasive||Gopal, 1987; Kairo et al., 2003; EPPO, 2014|
|Saint Lucia||Present||Introduced||Graveson, 2012||Naturalized|
|Saint Vincent and the Grenadines||Present||Introduced||Gopal, 1987|
|Trinidad and Tobago||Present||Introduced||Gopal, 1987|
|United States Virgin Islands||Present||Introduced||Acevedo-Rodríguez and Strong, 2012|
|Argentina||Restricted distribution||Gopal, 1987|
|Bolivia||Restricted distribution||Gopal, 1987|
|Brazil||Widespread||Native||Gopal, 1987; EPPO, 2014|
|-Espirito Santo||Present||Native||Lorenzi, 1982|
|-Mato Grosso||Present||Native||Gopal, 1987|
|-Mato Grosso do Sul||Present||Native||Lorenzi, 1982|
|-Minas Gerais||Present||Native||Gopal, 1987|
|-Rio de Janeiro||Present||Native||Gopal, 1987|
|-Rio Grande do Norte||Present||Native||Lorenzi, 1982|
|-Rio Grande do Sul||Present||Native||Gopal, 1987|
|-Santa Catarina||Present||Native||Gopal, 1987|
|-Sao Paulo||Present||Native||Gopal, 1987|
|Chile||Restricted distribution||Gopal, 1987; EPPO, 2014|
|Colombia||Widespread||Native||Invasive||Gopal, 1987; PIER, 2013; EPPO, 2014|
|Ecuador||Restricted distribution||Gopal, 1987; EPPO, 2014|
|Peru||Restricted distribution||Gopal, 1987; EPPO, 2014|
|Suriname||Restricted distribution||Gopal, 1987|
|Venezuela||Present||Gopal, 1987; EPPO, 2014|
|Belgium||Introduced, not established||Introduced||DAISIE, 2013|
|Czech Republic||Restricted distribution||Introduced||Not invasive||Pysek et al., 2002|
|France||Restricted distribution||Introduced||Not invasive||Georges and Pax, 2002; EPPO, 2014|
|-Corsica||Transient: actionable, under eradication||DAISIE, 2013; EPPO, 2014|
|Hungary||Introduced, not established||Introduced||DAISIE, 2013|
|Italy||Present||Introduced||DAISIE, 2013; EPPO, 2014|
|-Sardinia||Present, few occurrences||EPPO, 2014|
|-Sicily||Introduced, not established||Introduced||DAISIE, 2013|
|Portugal||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Russian Federation||Absent, formerly present||EPPO, 2014|
|-Central Russia||Absent, formerly present||EPPO, 2014|
|American Samoa||Present||Introduced||Invasive||PIER, 2013; EPPO, 2014|
|Australia||Restricted distribution||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|-Australian Northern Territory||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|-New South Wales||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|-Queensland||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|-South Australia||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|-Victoria||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|-Western Australia||Restricted distribution||Introduced||Invasive||Gopal, 1987|
|Cook Islands||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Fiji||Widespread||Introduced||1905||Invasive||Parham, 1958; EPPO, 2014|
|French Polynesia||Present||PIER, 2013; EPPO, 2014|
|Guam||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|Marshall Islands||Present||EPPO, 2014|
|Micronesia, Federated states of||Present||EPPO, 2014|
|Nauru||Present||Introduced||Invasive||PIER, 2013; EPPO, 2014|
|New Caledonia||Present||Introduced||Invasive||PIER, 2013; EPPO, 2014|
|New Zealand||Restricted distribution||Introduced||Gopal, 1987; EPPO, 2014|
|Northern Mariana Islands||Present||EPPO, 2014|
|Papua New Guinea||Widespread||Introduced||Invasive||Schmedding, 1995; EPPO, 2014|
|Samoa||Restricted distribution||Introduced||Invasive||Space and Flynn, 2000; EPPO, 2014|
|Solomon Islands||Present||Introduced||Invasive||Gopal, 1987; EPPO, 2014|
|US Minor Outlying Islands||Present||EPPO, 2014|
|Vanuatu||Present||Introduced||Invasive||PIER, 2013; EPPO, 2014|
History of Introduction and SpreadTop of page
The origin of E. crassipes is almost certainly the Amazon basin of Brazil (Barrett and Forno, 1982), and the natural distribution prior to 1800 is not thought to have extended beyond South America. Although it may not be strictly native in Central America it had certainly spread to many countries of Central America and the Caribbean by the end of the nineteenth century. It was first introduced to the USA (Louisiana) in 1884, when the plant was distributed to participants in the New Orleans Cotton Exposition and apparently became a problem thereafter (Julien, 2001), and further to Florida in 1890.
Because of its striking flowers, it was deliberately introduced into botanic gardens in many other countries, from which it inevitably spread as a weed. Some dates of introduction indicated by Gopal (1987) include: Australia, Egypt and Japan all about 1890; Indonesia, 1894; India, 1896; China, 1902 (1901 according to Xie Yan et al., 2001); Singapore, 1903; Sri Lanka, 1904; South Africa, 1910; the Philippines, 1912; Myanmar, 1913. It was probably introduced to Madagascar around, or shortly after, 1900 as an ornamental and was first recorded in 1920 (Binggeli, 2003).
The early introduction and spread of the plant to South-East Asia has been outlined by Burkill (1935). It was brought from an unknown location to Java in 1894, to Tonkin in 1902, and in about 1902 it reached southern China. A person noted the species in Hong Kong, admired its beauty and took the plant to Sri Lanka. A Chinese resident in Singapore imported it from Hong Kong to his garden and the plant was subsequently brought into the Botanic Gardens. Then local Chinese villagers took the plant to their homes and successfully fed it to their pigs and it became generally adopted for this purpose.
Within South-East Asia, there has been extensive spread throughout Malaysia, Indonesia, the Philippines, Vietnam, Thailand, Cambodia and Laos; also through southern provinces of China and Japan. E. crassipes was first reported from Papua New Guinea in 1962.
In recent years, introduction (deliberate and otherwise) has been especially serious in Africa, with troublesome infestations developing in the Congo river from about 1950, the Sigi and Pangani rivers (Tanzania) from 1955, 1959 respectively (Ivens, 1989), the upper Nile from about 1956, Senegal from about 1960 (all cited in Gopal, 1987). It was first recorded in Sudan in 1957 and is thought to have been introduced that year or shortly before. It started to spread rapidly up the Nile's tributaries thanks to steamer traffic (Gay, 1960). It was recorded along the Shire river (Malawi) from 1968 (Harley, 1993), Nigeria from 1982 (Akinyemiju, 1987), Ghana from 1984 (de Graft-Johnson, 1993), Benin from 1985 (van Thielen, 1993), Lake Kyoga (Uganda) from 1988, Lake Naivasha (Kenya) from 1989 (Owiti, 1990), and Lake Victoria from 1989 (Twongo, 1993). Although E. crassipes was present in Uganda before 1987 in the (relatively) lower reaches of the White Nile (Gopal, 1987), it was only noticed in Lake Kyoga by 1988 (Twongo, 1993). Occurrence in the Niger river in Mali and Niger has now also been confirmed (Lomer, 1995).
Once introduced to favourable habitats, especially open waters, E. crassipes may spread very rapidly and can form dense monotypic mats. In the 1950s, within 3 years of its first sighting, it had spread 1600 km along the Congo River (Holm et al., 1969). On Lake Victoria the species-spread in the early 1990s was just as dramatic but by the end of the decade the population had crashed (Mailu, 2001). In Madagascar, the potential threat to the freshwater bodies of the island was recognised in the 1920s following the introduction of the species as an ornamental. However, the advice to eradicate the plant was not heeded and by the late 20th century it became a major pest (Binggeli, 2003).
Risk of IntroductionTop of page
From the early part of the twentieth century E. crassipes has been identified as a troublesome plant and declared a noxious weed. For instance, in Fiji it was proclaimed a noxious plant, it was one of the first plants to be recognised as a noxious weed in January 1923 and growing it in a lily pond was made illegal (Parham, 1958). E. crassipes is also listed as a noxious weed in other countries, including Australia and South Africa. Even the movement of plant material may be prohibited within countries such as Australia (Parsons and Cuthbertson, 1992). There may also be regulations requiring the destruction of E. crassipes wherever it is found. However, enforcement is difficult and uneven. The lack of effective regulatory control has been responsible for most of the world's worst infestations of E. crassipes. The current ease with which plants, including E. crassipes, are available from the internet threatens efforts to prevent the sale and spread of weedy species to countries blacklisting them.
HabitatTop of page
E. crassipes is a floating weed of tropical and sub-tropical freshwater lakes and rivers, especially those enriched with plant nutrients. It may also be a weed in flooded rice.
Habitat ListTop of page
|Terrestrial – Managed||Cultivated / agricultural land||Present, no further details||Harmful (pest or invasive)|
|Freshwater||Present, no further details||Harmful (pest or invasive)|
Host Plants and Other Plants AffectedTop of page
|Oryza sativa (rice)||Poaceae||Main|
Biology and EcologyTop of page
The chromosome number is 2n = 32 (Darlington and Wylie, 1955).
Physiology and Phenology
The growth of E. crassipes is extremely rapid and the plant may double its population size in 6 to 18 days.
E. crassipes leaves show anatomical characteristics of C3 plants but the photosynthetic process shows some characteristics of the more productive C4 plants, especially in showing no light saturation up to high light levels. This makes it a highly efficient plant and relative growth rates have been recorded of 1.012 to 1.077. Other studies suggest it is capable of increasing in biomass by up to 12% per day. The time required to double in number or biomass is variously reported to be from 6 to 15 days. Productivity can also be expressed in terms of 100-500 g fresh weight/m² per day, 1000-5000 kg/ha per day or 400-1700 t/ha per year. The total biomass or 'standing crop' can be as much as 42 kg/m² or 420 t fresh weight/ha. As the dry weight is normally about 5-7% of fresh weight, this represents about 2.5 kg dry weight/m² or 25 t dry matter/ha (Gopal, 1987). As would be expected, the foliage is very dense with one study in Florida finding leaf area index values of 7.8 and 5.8, comparable with many of the most productive terrestrial ecosystems (Knipling et al., 1970).
Flowering is seasonal in some countries but not obviously so in others. There is apparently little or no response to photoperiod but considerable evidence that flowering may be induced by nutrient shortages.
The flowers of E. crassipes are tristylous, but unlike some other tristylous species, there is no incompatibility between the different forms. Hence pollination (mainly by wind) can result in good seed set, though in some populations there may be a higher degree of self-incompatibility.
E. crassipes propagates vegetatively and by seed. After flowering, the peduncle is deflexed and the capsules mature and seeds are eventually released below water. The seeds are capable of germinating immediately but may remain dormant for many years. Germination is encouraged by aerobic conditions and alternating temperatures; large populations of seedlings may become established on exposed mud at the edges of water bodies when water levels fall. Seedlings are rooted in mud initially but become free-floating as a result of wave action or rising water levels. From an early stage, the axillary buds of the older leaves of the seedling are capable of developing into stolons, which grow horizontally and develop daughter plants. Such vegetative spread can occur indefinitely and very large populations are produced in this way without any sexual reproduction.
Optimum temperature for growth of E. crassipes is 25-30°C. Growth ceases when water temperature is above 40°C or below 10°C, but short periods at freezing may be tolerated.
E. crassipes is very responsive to nutrients (especially nitrogen and phosphorus) and high growth rates are always associated with eutrophic, nutrient-rich conditions. Growth rate was greater by a factor of eight where total nutrient content was 52 mg/l, compared with 8 mg/l (Lugo et al., 1979). The growth rate is proportional to the percentage concentration of nitrogen in the leaves (Aoyama and Nishizaki, 1993) and there is a hyperbolic relationship between the growth rate and the nutrient concentration in the water. The mean percentage nitrogen (dry weight) of the second youngest leaves in the field has been determined at between 1 and 5% (Center and Wright, 1991; M Purcell, unpublished data; MH Julien, unpublished data; Lorber et al., 1984). The percentage phosphorus is slightly lower - ranging from 0.2 to 1.0% (cited in Lorber et al., 1984). The percentage nitrogen varies between plant parts and decreases exponentially as leaves age (Center and Wright, 1991).
Optimum pH is between 6 and 8 and extremes of pH (below 4.5 or above 10) can be damaging. Calcium concentration is important, with an observed threshold of 5 mg/l, below which growth ceases.
E. crassipes will tolerate only low levels of salinity; one-quarter strength sea water is lethal (Muramoto et al., 1991) and the problem in coastal lagoons depends on the growth of the weed in the fresh water of the rivers that flow into the lagoon.
E. crassipes is often associated with other water weeds such as Pistia stratiotes, Myriophyllum aquaticum and Azolla filiculoides. However, it tends to be the dominant species unless some form of biological control has been initiated (Chikwenhere, 2001).
Air TemperatureTop of page
|Parameter||Lower limit||Upper limit|
|Absolute minimum temperature (ºC)||0|
Soil TolerancesTop of page
Water TolerancesTop of page
|Parameter||Minimum Value||Maximum Value||Typical Value||Status||Life Stage||Notes|
|Salinity (part per thousand)||0||6||Optimum|
|Salinity (part per thousand)||8||Harmful|
|Water pH (pH)||6||8||Optimum|
|Water temperature (ºC temperature)||10||25||Optimum|
|Water temperature (ºC temperature)||5||Harmful|
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Flechtmannia eichhorniae||Herbivore||Whole plant|
|Neochetina bruchi||Herbivore||Growing point/Leaves/Stems||Africa; Benin; India; New South Wales; Queensland; Papua New Guinea; Zimbabwe|
|Neochetina eichhorniae||Herbivore||Growing point/Leaves/Stems||Africa; Benin; India; Malaysia; Pacific Islands; Queensland; South Africa; Sri Lanka; Texas; Thailand; New South Wales; Papua New Guinea; Indonesia; Zimbabwe|
|Niphograpta albiguttalis||Herbivore||Growing point/Leaves/Stems||Africa; Queensland; New South Wales|
|Trichechus manatus||Herbivore||Whole plant|
Notes on Natural EnemiesTop of page
Almost 100 different insect species and a comparable number of pathogens have been recorded as attacking E. crassipes (refer to Gopal, 1987). Most of these are restricted to the areas of the New World from which the weed originates. In Africa and Asia, the weed is normally quite healthy, though sporadically attacked and sometimes moderately damaged by sundry local organisms. A few species of insects and fungi have been developed for use as biological control agents, with varying success (see Control).
The more important natural enemies in South America are listed, including those that have been used as biological control agents, or studied as potential biological control agents, and also some organisms that have been reported as causing significant damage in some of the countries where E. crassipes has been introduced (see Waterhouse (1987), Julien and Griffiths, (1998) and Hill et al. (1999) for details).
Means of Movement and DispersalTop of page Natural Dispersal (Non-Biotic)
Wind will readily move the plant and the upright leaves act as sails in lakes and canals. Along rivers, water flow is the prime mover of vegetative material but strong winds may sometimes blow the plant upstream.
Vector Transmission (Biotic)
Seeds are thought to be transported over long distances by birds (e.g. waterfowl and shore birds) and if coated in mud they may cling to both mammals and birds (Holm et al., 1969; Batcher, 2000).
New infestations may arise via unintentional human transportation such as canoes, boats and probably even charcoal transport as sacks used in the process are, in some parts of Africa, plugged with the plant.
The high ornamental value of the plant still makes it liable to intentional introductions, especially as the species is up for sale on the internet.
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Economic ImpactTop of page
As a result of its rapid growth and large biomass, E. crassipes has a range of detrimental effects, which include:
- Physical interference with water transport, communication and access. Gopal (1987) refers to serious interference with navigation in southern USA, South Africa, southeast Asia, Australia, Congo and Sudan. Annual costs of control or removal have, in the past, amounted to millions of dollars on the Panama Canal, on the Nile in Sudan, on the Congo and have been as much as $35 million in southern USA. Costs of controlling water hyacinth in Malaysia have been estimated at M$ 10 million per year (Mahomed et al., 1992), while Harley et al. (1996) quoting this figure, state that present actual costs are believed to be much higher. In recent years, the operation of Port Bell, Uganda, on Lake Victoria has been seriously threatened and costs have involved $1 million for a mechanical harvester, as well as the loss of trade at times when the port was completely blocked (Hill, 1999). Infestations are also increasing in Ethiopia, creating a range of problems including restricted access (Aweke, 1994). Harley et al. (1996) refer to 'devastating effects' on socio-economic structure and on the environment in the lower flood plain of the Sepik river in Papua New Guinea resulting from problems of access to subsistence gardens, hunting and fishing areas, and markets. The same authors refer to the recent increase in water hyacinth infestations in West Africa which are resulting in serious disruption of the socio-economic structure, food supply and health of several million people. In Nigeria, Alimi and Akinyemiju (1991) showed that costs of fuel and repairs to boats on infested waterways was approximately three times that on uninfested waterways. The problem has also been increasing recently in Mali (Dembele et al., 2000). Economic losses also result from interference with recreational uses of water bodies (for example, Gopal, 1987; Aweke, 1994; Cilliers et al., 1996).
- Interference with fishing. This effect is most acute for small-scale fishing communities. Apart from the problems of access to fishing grounds and interference with the spreading or retrieval of nets or with landing their catch, there can be serious effects on fish stocks and fish breeding. Although a sparse cover of water hyacinth may not reduce fish and may even be used to advantage in some fishing techniques (Gopal, 1987), a dense infestation can lead to de-oxygenation and kill-off fish or reduce fish stocks. Gopal (1987) refers to heavy losses of fish production in the Congo, Nile and other rivers and in Pakistan and to losses amounting to 45 million kg in West Bengal, India in the 1950s and reductions of 70% in fish production in the USA as a result of a cover of only 25%, presumably due to reduction of phosphorus levels and phytoplankton. The shallow water of lake edges can be especially important spawning areas for fish and a dense cover of water hyacinth can interfere severely with fish breeding. Hill (1999) refers to this phenomenon on Lake Victoria where the estimated 10,000 ha of the weed includes an almost continuous fringe along the shoreline extending to at least 10 m. Labrada (1996) quotes fuel costs increased by a factor of 2-3 and fish catches down 50-75% on parts of Lake Victoria. Fishermen affected by another relatively new infestation, in the Shire river in Malawi, report reduced catches which are not confirmed by the locally available statistics but there is no doubt fishermen are being troubled by a reduced range of fish species, loss of nets and impeded access (Terry, 1996).
- Risks of mechanical damage to hydro-electric installations and other structures such as bridges. Expensive barriers or mechanical harvesters may be needed to minimize these risks, for example, to the Owen Falls Dam on Lake Victoria (Hill, 1999). Elsewhere, there are similar concerns in South Africa (Harley et al., 1996), Brazil (Pitelli, 2000), New Zealand (Clayton, 2000) and Ethiopia (Aweke, 1994).
- Reduced irrigation flow can indirectly cause crop loss but there can also be direct interference and competition from water hyacinth where it occurs in flooded rice. Such losses have been estimated at many million dollars in West Bengal, India and as significant in many other countries including Sri Lanka, Bangladesh, Burma, Malaysia, Indonesia, Thailand, Philippines, Japan and Portugal (Gopal, 1987).
Nang’alelwa (2008) summarizes the socioeconomic effects in the Victoria Falls World Heritage site in Zambia. Major impacts include effects on the generation of hydro electric power, tourism development, native biodiversity, fish catches and human health. Other recorded impacts are reduced quality and quantity of water for domestic use, restricted navigation of waterways and the threat posed to vital infrastructure.
Environmental ImpactTop of page
Once it proliferates in a water body, E. crassipes dramatically alters the ecosystem and often results in environmental degradation and a reduction in bio-diversity. A number of authors note that in many water bodies and wetland areas, the encroachment of water hyacinth has reduced or eliminated natural vegetation (Terry, 1996; Kumar and Rohatgi, 1999). The plant may negatively impact some native species of invertebrates, fish, birds and plants. For example, in Madagascar, many parts of the Alaotra Lake, a site of biological importance, have been reported as covered with carpets of E. crassipes that are detrimental to a number of species, such as the duck Thalassornis leuconotus (Binggeli, 2003).
Other environmental impacts include:
- Restricting water flow in rivers, irrigation and drainage channels, thus reducing irrigation water and/or leading to greater risk of flooding. Gopal (1987) refers to water flow being reduced by 40-95% in irrigation channels, sometimes leading to flooding in Malaysia and Guyana.
- Excess evapotranspiration, causing wastage of water that would otherwise be used for irrigation, drinking, fisheries, etc. Rates of loss have been reported up to 13 times that from a free water surface, with an average of 2.5 times the loss (Gopal, 1987). In India, the loss of water of the mats of E. crassipes was 7.8 times greater that of open water thus resulting in massive wastage of water especially in dry regions (Vasudevan and Jain, 1991). However, it has recently been claimed that these figures have been grossly exaggerated by inadequate experimental technique (Allen et al., 1997).
- When mats decompose dissolved oxygen levels are reduced and sedimentation increases.
The effects of E. crassipes on physicochemical characteristics of water in Lake Naivasha, Kenya, are described by Mironga et al. (2012). Impacts include greater levels of free carbon dioxide, lower pH and lower levels of dissolved oxygen in infested areas than in open water. A similar study in Badagry Creek and Ologe Lagoon, Lagos, Nigeria (Ndimele, 2012) found effects on salinity, conductivity, total hardness and total dissolved solids. It is suggested that while there are negative impacts on water quality, the ability of E. crassipes to passively absorb heavy metals and nutrients can be put into good use.
Threatened SpeciesTop of page
|Threatened Species||Conservation Status||Where Threatened||Mechanism||References||Notes|
|Rostrhamus sociabilis plumbeus (Everglade snail kite)||USA ESA listing as endangered species USA ESA listing as endangered species||Florida||Ecosystem change / habitat alteration||US Fish and Wildlife Service, 2008|
Social ImpactTop of page
E. crassipes may reduce water quality in various ways and encourage mosquitoes, snails and other organisms associated with human illnesses, including malaria, schistosomiasis, encephalitis, filariasis and cholera (Gopal, 1987). Harley et al. (1996) comment that people in Papua New Guinea have died through a combination of reduced nutrition, degraded water, increased disease vectors and generally reduced health, directly related to the degrading effect of water hyacinth on the environment. Dense mats greatly hinder boating by fishermen and may prevent fishing altogether, thus denying the locals their main source of protein and sometimes forcing people to relocate. In extreme cases of competition between E. crassipes and rice crops, fields have been abandoned. In the Lake Victoria Basin, the main negative social impact were identified by interviewees as an increase in certain diseases, difficulties associated with clean water availability and migration of communities (Mailu, 2001).
Risk and Impact FactorsTop of page Invasiveness
- Invasive in its native range
- Proved invasive outside its native range
- Highly adaptable to different environments
- Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
- Highly mobile locally
- Has high reproductive potential
- Has propagules that can remain viable for more than one year
- Damaged ecosystem services
- Ecosystem change/ habitat alteration
- Negatively impacts agriculture
- Negatively impacts tourism
- Reduced amenity values
- Reduced native biodiversity
- Competition - monopolizing resources
- Highly likely to be transported internationally deliberately
- Difficult to identify/detect as a commodity contaminant
- Difficult/costly to control
UsesTop of page
E. crassipes can be utilized in various ways (for instance for East Africa see Lindsey and Hirt, 1999). Although not generally suitable as an animal feed, small amounts can be fed to pigs and buffaloes, but in China during the 1950s-1970s, when fodder was scarce, it was widely used as an animal feed (Ding Jianqing et al., 2001). It can be used as a mulch, for making compost, fuel bricks, paper or board, for generating methane biogas, and for removing nutrients and toxic chemicals from water. Recent work on composting includes Montoya et al. (2013) who found that a large-scale composting system using water hyacinth as a primary feedstock reached high enough temperatures to inactivate seeds and other propagules, and thus that the plant can be composted without the potential danger of spread.
Its very high growth rate and ability to withstand various types of pollution are proving of interest for the treatment of polluted water but there remains the problem of disposal of the harvested (polluted) material (Aoyama et al., 1986; Ayade, 1998). Yan et al. (2012) tested E. crassipes for removal of pollutants in Lake Caohai, China, and found that the plant could not only remove phosphorus in the water, but also remove the soluble phosphorus in the sediment of Lake Caohai, Ndimele and Ndimele (2013) suggest that the species absorbs petroleum hydrocarbon and can be used for phytoremediation of crude oil-polluted aquatic ecosystems.
Potentially, water hyacinth could be very important in sewage and waste water treatment. Its fast growth rate and high absorption of nutrients and heavy metals could make it a cheap and largely environmentally benign form of decontamination (Hill et al., 1999; Zhu et al., 1999). However, the biggest use made of water hyacinth is probably as an ornamental in temperate regions (Cohen, 1993).
Work on utilization includes use as an organic manure in Bangladesh (Nasima et al., 1997); as a compost to suppress nematodes in India (Verma et al., 1997); for water purification (Ayade, 1998); for biogas production (Rodriguez et al., 1997; Sarkar and Banergee, 2013)); for feeding buffaloes in India (Mitra et al., 1997); and as a mulch to suppress weeds in Indonesia (Lamid and Wahab, 1996). Masto et al. (2013) explored the conversion of E. crassipes to biochar for improvement of soil quality. There are many recent studies on utilizing E. crassipes for bioenergy. Hussain et al. (2013) converted E. crassipes biomass into liquid hydrocarbon fuel using catalytic pyrolysis. Bergier et al. (2012) suggest that biomass from water hyacinth in the Panatanal of South America could be managed for production of biofuels. Sudhakar et al. (2013) assess bioelectricity production using water hyacinth biomass. Anaerobic co-digestion with poultry litter for biogas production is considered by Patil et al. (2013), while Zhang et al. (2013) report on hydrothermal liquefaction. Biogas production from water hyacinth polluting water bodies in Nigeria is studied by Adeleye et al. (2013).
Uses ListTop of page
Animal feed, fodder, forage
- Fodder/animal feed
- Soil improvement
Detection and InspectionTop of page
Detection of mature floating E. crassipes plants is all too simple but where control methods have been used to eliminate these, there is a need to watch for seedling plants at the edges of the water body.
Similarities to Other Species/ConditionsTop of page
Apart from the possible confusion with other species of Eichhornia, notably E. azurea in Central and South America (see Taxonomy and Nomenclature), some other species in Pontederiaceae could perhaps be confused with E. crassipes. These include the rice weed Monochoria vaginalis which is common throughout South-East Asia. This is superficially similar, with a spike of showy purple flowers, but these are smaller, radially symmetrical and the petals are free. Leaf shape can be somewhat similar but petioles are not swollen. Some species of Pontederia can occur as aquatic weeds in North America but these have a unilocular ovary and flowers are two-lipped, each lip with three lobes. Several species of Heteranthera occur as weeds in the Americas and Africa but these have only three stamens. All these related species are rooted weeds, not floating aquatics like E. crassipes.
Prevention and ControlTop of page
Although the exact nutrient threshold below which E. crassipes will not flourish is not yet clear, it is certain that its vigour is directly related to available levels of nitrogen and phosphorus. Wherever possible, nutrient levels in the water body should be reduced or controlled: for example, by processing sewage or other nutrient-rich water, or by diverting it away from critical areas In South Africa, Coetzee and Hill (2012) suggest that the first step in any control programme should be to reduce the nutrient status of the water body, as a meta-analysis of studies that investigated the combined effect of P and nitrogen (N) water nutrient concentration and control agent herbivory showed that water nutrient status was more important than herbivory in water hyacinth growth.
Where E. crassipes is causing the most acute problems (e.g. impeding access for fishermen, or threatening to block harbours or damage hydro-electric installations), an effective solution may be the use of floating booms or fixed barriers to prevent movement into the critical areas. Booms may also be used to try and prevent movement of the weed down rivers, though their success will depend on their design (complicated by the need to maintain navigability along the river), the mass of material involved and the capacity to clear the booms by physical removal of weed.
Physical removal or destruction of the infestation may be achieved on a small scale by manual removal. On the larger scale, machinery is needed, either shore-based, or mounted on boats. Where possible, on smaller water bodies, reliance should be placed on unspecialized shore-based equipment (e.g. drag-lines, excavators, moving-belt elevators etc.), the weed being pushed to the shore by suitably modified boats. For larger water bodies, special boats may be needed with suitable harvesting equipment, together with a means of crushing the weed or otherwise reducing the volume of water. Where the water body is sufficiently large and deep for the weed to be returned to the water after crushing, without risk of decomposition causing deoxygenation, the use of such equipment may be economic. If the weed has to be transported to the land for unloading, the running costs become much greater and such methods may not be economic.
Julien (2008) reviews biological aspects of E. crassipes related to management, and suggest that containment and eradication from a catchment may only be accomplished if the invasion is very young, small, isolated and accessible, and if the short-term resource commitment is high. Jyoti and Garima (2013) present methods of control including manual pulling and harvesting.
2,4-D has been widely used for control of E. crassipes. Best results are achieved under conditions of rapid growth, high temperature and high humidity, when most plants of any age will be killed and sink within 2-4 weeks. Under less favourable conditions, some plants may regrow and require repeat treatment. In any case re-treatment is almost inevitably required after a few months as a result of re-infestation from incompletely sprayed plants, re-invasion from outside the sprayed area, or regrowth by seedlings.
Glyphosate has been tested and used for control of E. crassipes. It is much more expensive than 2,4-D but has possible advantages over 2,4-D in not causing taint of drinking water and in causing a slower kill of the weed, apparently reducing the risks of deoxygenation during decomposition (Findlay and Jones, 1996).
Other herbicides that have been used include the contact herbicides paraquat and diquat, but these have high mammalian toxicity and should not normally be used. Diquat use is described by Pitelli et al. (2011), who suggest that night application is more effective than day spraying. Aminotriazol [amitrole], ametryn and terbutryn can each be effective alone, but have been most often used in mixture with 2,4-D. Wersal and Madsen (2010) evaluated the use of penoxsulam, which gave effective control which was not improved by applying in combination with diquat.
New herbicides in the imidazolinone and sulfonylurea groups have been shown to have high activity on E. crassipes, but have not yet been adequately tested. These and other possibilities have been summarized by Price (1993).
Herbicides have rarely been used with complete success, owing to the need for repeated treatment over a long period, requiring dedicated management and organization. Apart from the problems of limited success, the use of 2,4-D and other herbicides can be unsatisfactory in several other respects. Ester formulations of 2,4-D can be highly toxic to aquatic organisms as well as creating a vapour drift problem. While the direct toxicity to aquatic organisms of 2,4-D amine salt formulations and the other listed compounds is largely negligible at the concentrations reached in the water, there can be devastating stress caused by deoxygenation as the weed dies and decomposes. Other problems include those of taint of drinking water by 2,4-D, and, for any herbicide that is used, damage by spray-drift onto non-target crops and other plant life adjacent to sprayed areas.
Seven arthropods and three fungi have been developed and released for the biocontrol of E. crassipes (Harley, 1990; Julien and Griffiths, 1998). The arthropods are the curculionids Neochetina bruchi and Neochetina eichhorniae, the pyralids Xubida infusellus and Niphograpta albiguttalis, the noctuid Bellura densa, the mirid Eccritotarsus catariensis, and the galumnid mite Orthogalumna terebrantis. The fungi are all hyphomycetes: Acremonium zonatum, Cercospora piaropi and Cercospora rodmanii. Additionally, there has been work on the development of the fungus Alternaria eichhorniae as a mycoherbicide (Aneja, 1996; Shabana, 1997). Acremonium zonatum, Cercospora piaropi, Myrothecium roridum, and Rhizoctonia solani are viewed as suitable bioherbicides (Charudattan, 2001). In Africa, an international programme has been established to develop a mycoherbicide for the control of the weed, using fungal isolates that have been found in Africa (Bateman, 2001). Karim Dagno et al. (2012) review the current status of development of mycoherbicides against E. crassipes, but report that biological, technological and commercial constraints have hindered progress. Oil emulsions are recognized as a way to increase both efficiency of application and efficacy of biocontrol agents
The two Neochetina weevils have together given excellent results in the USA, Argentina, India, Australia and Sudan, acting apparently in a complementary fashion. Infestations of E. crassipes have been reduced by 80-90% or more. In Uganda, the two weevils have greatly reduced the problem on Lake Kyoga, and are beginning to take effect on Lake Victoria (Hill, 1999). In Papua New Guinea, N. eichhorniae is reported to be giving 'permanent control' in some areas (Orapa and Atip, 1996). More recently, Orapa and Julien (2001) reported that although control had been achieved in some areas, such as the Sepik River and Waigani Lake, the full impact of biological control by the Neochetina weevils on water hyacinth in PNG is not known.
Some successful control programmes have been recorded in Mexico (Panduro and Domunguez, 1998), Benin, South Africa, Zimbabwe and Malawi. Control takes from 2 to 10 years depending on the location and the environmental conditions, but in some locations (including the countries mentioned) the weevils do not appear able to control the weed.
Adult weevils feed on the leaf and petiole surfaces, preferentially on the youngest leaves (Center, 1985). They make distinctive, almost square, feeding scars. This may cause significant loss of functional leaf surface and also may allow entry of pathogens, with the potential in extreme situations for removing over 50% of the laminar area (Van and Center, 1994). However, the most significant damage is caused by the larval stages. Eggs are laid in the petioles. Upon hatching, the larvae burrow down the petiole into the crown of the plant where they can cause major damage (Patnaik et al., 1988). The weevils pupate underwater in the roots. Under certain circumstances the adults can migrate through flight (Buckingham and Passoa, 1984). This damage to the petiole often results in complete collapse of the leaf and eventually in loss of buoyancy so that the whole plant sinks. Each of the two Neochetina species has small but distinct differences in biology, ecology and feeding habits, which result in additive, complementary effects. N. bruchi are slightly smaller weevils and develop faster but in many locations including Florida, USA, and Benin, N. eichhorniae is the species most commonly encountered in the field. The developmental time is much shorter in the tropics with N. eichhorniae taking 80 days to develop from egg to adult in Florida and about 50 days in West Africa.
The moth Niphograpta albiguttalis is believed to have contributed to the successes in Sudan and the USA. Oke et al. (2012) report that this moth did not successfully establish when released in Benin or Ghana, but that without recorded release of the moth in Nigeria the larvae were found damaging water hyacinth in the infested waterways of Badagry, Ejirin and Epe in Lagos State and Iwopin in Ogun State. The larval instars found were damaging only water hyacinth with bulbous petioles. The other organisms listed above have rarely been effective on their own, but the fungi are often observed to increase the damage caused by insects or by the mite Orthogalumna terebrantis; this has been observed in South Africa.
Chemical control (e.g. using 2,4-D) may be necessary as an extreme measure, for the rapid destruction of large masses of weed which are seriously impeding access or navigation. All the larvae of Neochetina spp. and many adults on the sprayed plants are likely to be lost as a result of complete kill of the weed. This should be considered in deciding the areas to be treated, in addition to the possible problems from deoxygenation when the weed is decomposing. Where Neochetina spp. are being introduced, any herbicide treatment should of course be kept well away from the introduction points. Low doses of 2,4-D, which damage but do not kill the weed are believed to encourage insect attack and will thus be beneficial in the longer term (Haag and Habeck, 1991). Other evidence suggesting that herbicides are not necessarily detrimental to Neochetina spp. is provided by, for example, Findlay and Jones (1996) and Center et al. (1999). Herbicides are also known to encourage certain fungi. Hence chemical and biological control are not necessarily incompatible.
Biological control programmes can readily involve local community groups. In Australia, CSIRO has harnessed the resources of the school system via the formation of the Double Helix Science Club as part of a sponsored initiative to promote science in schools. In 1995, this club released the biocontrol agent Neochetina bruchi (Briese and McLaren, 1997).
A new agent, Cornops aquaticum, is being tested for specificity in South Africa (Oberholzer and Hill, 2001). Coetzee et al. (2011) review biological control efforts in South Africa, but suggest that long-term management of alien aquatic plants in South Africa relies on the prevention of new introductions of aquatic plant species that could replace those that have been controlled, and, more importantly, on a reduction in nutrient levels in South Africa's aquatic ecosystems.
Sacco et al. (2013) evaluate the potential of the planthopper Taosa logula, native to South America, for control of E. crassipes. Tests showed that individual growth and biomass production of water hyacinth was reduced due to the effect of the insect feeding above five nymphs per cage. The number of new plants produced by clonal reproduction was only significantly different above 15 nymphs per cage. These results suggest that this planthopper could be an effective agent for the biological control of E. crasssipes.
Although it is hoped that biological control will eventually be capable of achieving the necessary level of control of E. crassipes, there is likely to be scope for the integration of physical and chemical methods with biological methods on a local basis, to help speed the achievement of control. The possible approaches include:
- control of nutrient levels.
- use of booms to control movement of the weed.
- exploitation of variable water levels.
- manual removal of the weed from shores and small channels.
- mechanical removal or destruction by land-based or floating equipment.
- use of biological control agents.
- careful use of herbicide to kill or weaken the weed.
- utilization of the weed.
An example of a well integrated control approach (in Mexico) is provided by Gutierrez et al. (1996). In South Africa, biological control with five arthropod species and fungal pathogens attempted since the mid-1970s has had limited success and it has been suggested that additional control agents may be required as well as implementing site-specific integrated management plans (Hill and Cilliers, 1999). Due to the weed's recent rapid increase in the species' abundance and distribution in Africa and elsewhere, international co-operation has been promoted in order to effectively combat the plant (Julien et al., 1996). Lu et al. (2007) suggest that in China the currently dominant biological control-centered view should be broadened to a sustainability science-based management framework that explicitly incorporates principles from landscape ecology and Integrated Pest Management.
Control of Nutrient Levels
The reduction of nutrient pollution of water bodies, wherever it is at all feasible, should be a high-priority approach. Redistribution of excess nutrient, as an alternative to its prevention, should be considered in some situations.
Where infestations occur in relatively narrow rivers, the removal by manual or land-based machinery is often feasible and, although such removal is expensive, the cost may be at least partly offset by utilization (see below). In larger water bodies, the weed should, wherever possible, be pushed to the shore for harvesting by land-based methods, but floating equipment may be appropriate in some situations.
A range of uses for water hyacinth have been proposed and studied (see Uses) none can be regarded as suitable for large-scale use and at the same time provide a satisfactory means of control. However, some of the uses can be exploited on a small scale, especially in conjunction with manual or mechanical harvesting, to recoup some costs and help to make the procedures more economic. Some of these can help to cover some of the costs of control but in almost no case does the usefulness outweigh the economic problems caused by the weed. The possibilities of incorporating utilization into an integrated system of control are reviewed in detail by Gopal (1987).
Each water body should be considered separately; an ideal combination of measures should be devised for each water body, depending on many factors and in close consultation with all users of the water.
Gopal (1987) ends his book with the warning that 'The interests of mankind can only be safeguarded by seeking effective control of water hyacinth and not by its utilization'.
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ContributorsTop of page
07/07/13 Updated by:
Julissa Rojas-Sandoval, Department of Botany-Smithsonian NMNH, Washington DC, USA
Pedro Acevedo-Rodríguez, Department of Botany-Smithsonian NMNH, Washington DC, USA
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