Azolla filiculoides (water fern)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Plant Type
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Host Plants and Other Plants Affected
- Growth Stages
- Biology and Ecology
- Latitude/Altitude Ranges
- Air Temperature
- Rainfall Regime
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Pathway Vectors
- Plant Trade
- Impact Summary
- Economic Impact
- Environmental Impact
- Threatened Species
- Social Impact
- Risk and Impact Factors
- Similarities to Other Species/Conditions
- Prevention and Control
- Gaps in Knowledge/Research Needs
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Azolla filiculoides Lamarck
Preferred Common Name
- water fern
Other Scientific Names
- Azolla bonariensis Bertoloni
- Azolla japonica Franch. & Sav.
- Azolla magellanica Willd.
- Azolla squamosa Molina
International Common Names
- English: fairy moss; mosquito fern; Pacific azolla; red water fern; water fern; water velvet
- Spanish: helecho acuatico; hlechito del aqua; lenteja de agua
- French: fougere d'eau
Local Common Names
- China: xi lu-ping; xi man jiang hong
- Germany: grosser algenfarn
- Japan: akaukikusa
- Netherlands: grote Kroosvaren
- South Africa: rooivaring
- AZOFI (Azolla filiculoides)
- AZOJA (Azolla japonica)
Summary of InvasivenessTop of page
A. filiculoides is a small fern native to the Americas which has spread widely throughout the world by a variety of mechanisms, of which man has become the most significant (Lumpkin and Plucknett, 1982). Man has introduced A. filiculoides into Europe, North and sub-Saharan Africa, China, Japan, New Zealand, Australia, the Caribbean and Hawaii. In eutrophic water systems, A. filiculoides grows rapidly, easily outcompeting indigenous vegetation. Decaying root and leaf matter below a mat of A. filiculoides, coupled with the lack of light penetration, creates an anaerobic environment which can reduce the quality of drinking water and make survival for other organisms in the water impossible.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Plantae
- Phylum: Pteridophyta
- Class: Filicopsida
- Family: Azollaceae
- Genus: Azolla
- Species: Azolla filiculoides
Notes on Taxonomy and NomenclatureTop of page
The origin of the generic name Azolla is thought to be derived from a conjugation of two Greek words meaning 'to dry' and 'to kill', inferring that the fern is killed by drought (Moore, 1969; Lumpkin and Plucknett, 1980).
The genus Azolla has no generally accepted taxonomic framework, primarily as a result of the plant's diminutive structure, morphological and phenotypic plasticity (Stergianou and Fowler, 1990; Saunders and Fowler, 1992) resulting in numerous synonyms. Currently, the genus is divided into three sections (Saunders and Fowler, 1992; Saunders and Fowler, 1993): section Azolla, which includes A. filiculoides Lam., A. rubra R. Br. (now more commonly regarded as a subspecies of A. filiculoides), A. caroliniana Willd., A. microphylla auct. non Kaulf, A. mexicana Presl and A. cristata Kaulf.; section Rhizosperma, which includes A. pinnata R. Br. var. africana (Desv.) R.M.K. Saunders and K. Fowler, stat. et comb. nov., A. pinnata R. Br. var. asiatica R.M.K. Saunders and K. Fowler, subsp. nov., and A. pinnata R. Br. var. pinnata; and section Tetrasporocarpia, which includes A. nilotica Decne. ex Mett. Taxonomic problems have primarily centred around the closely related species in section Azolla (Nayak and Singh, 1989; Stergianou and Fowler, 1990). A study of specimens by Evrard and Van Hove (2004) concluded that only two species exist in America and, according to the priority rule for nomenclature, they should be named A. filiculoides and A. cristata.
DescriptionTop of page
A. filiculoides is a small aquatic heterosporous fern, rarely larger than 25 mm (O'Keeffe, 1986). The genus is unique in that it grows in association with a heterocystous cyanobacterium (blue-green alga), Anabaena azollae Strasburger (Nostocales: Nostocaceae), which is located in cavities in the dorsal leaf-lobes (Ashton and Walmsley, 1984). This symbiotic association is the only one known between a pteridophyte and a cyanobacterium (Ashton and Walmsley, 1984). The Azolla macrophyte consists of a main rhizome, which branches into secondary rhizomes. These all bear alternately arranged small leaves. Ventrally, unbranched adventitious roots hang down into the water from nodes. Nutrients are absorbed directly from the water by the roots. In very shallow water, however, the roots may touch the soil thus deriving nutrients from it (Wagner, 1997). Rao (1936) reported that in A. pinnata, once the roots attain a length of 40-50 mm they drop off. Root hairs found along the length of the root provide accommodation for a large number of protozoa, algae and soil particles (Rao, 1936).
Plant TypeTop of page
DistributionTop of page
According to Lumpkin and Plucknett (1980), A. filiculoides is native to the Rocky Mountain states of the western USA and Canada, through Central America and to most of South America. It has been introduced to Europe, North and sub-Saharan Africa, China, Japan, New Zealand, Australia, the Caribbean and Hawaii.
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.Last updated: 30 Jun 2021
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Ghana||Present||Introduced||Invasive||Tano River and Lagoon, Kpong Headpond, Lower Volta River at Tefle, Sogakope, Ada; other water bodies countrywide; Original citation: deGraft-Johnson and Akpabey (2015)|
|South Africa||Present, Widespread||Introduced||1948||Invasive|
|Israel||Present, Few occurrences|
|Poland||Present, Few occurrences|
|Canada||Present||Present based on regional distribution.|
|Trinidad and Tobago||Present||Introduced||Original citation: Reed, 1965|
|-Rio Grande do Sul||Present||Native|
History of Introduction and SpreadTop of page
IntroductionsTop of page
|Introduced to||Introduced from||Year||Reason||Introduced by||Established in wild through||References||Notes|
|Natural reproduction||Continuous restocking|
|China||Germany||1977||Horticulture (pathway cause)||Yes||Lumpkin and Plucknett (1980)||Introduced for green fertiliser|
|Egypt||1980||Horticulture (pathway cause)||Yes||Searg et al. (2000)|
|Germany||1899||Yes||Hussner et al. (2010)|
|India||Horticulture (pathway cause)||Yes||Lumpkin and Plucknett (1980)||Introduced for green fertiliser|
|Iran||Horticulture (pathway cause)||Yes||Sadeghi et al. (2012)||Introduced for green fertiliser|
|Italy||Horticulture (pathway cause)||Yes||Introduced for green fertiliser|
|Mozambique||South Africa||2000||Yes||Langa (2013)||Natural dispersal|
|Poland||early 1900s||Aquaculture (pathway cause)||Yes||Szczesniak et al. (2009)|
|Portugal||1930||Horticulture (pathway cause)||Garcia-Murillo et al. (2007)||Introduced for green fertiliser|
|South Africa||Brazil||1948||Horticulture (pathway cause)||Yes||Hill (1999)|
|Spain||1957||Horticulture (pathway cause)||Yes||Garcia-Murillo et al. (2007)|
|UK||1920||Horticulture (pathway cause)||Yes||Janson (1921)|
|Zimbabwe||South Africa||1995||Yes||McConnachie et al. (2003)||Natural dispersal|
Risk of IntroductionTop of page
Further spread is likely as A. filiculoides continues to be sold in nurseries as a fish pond plant. In Africa and Europe, dispersal between countries will no doubt continue due to the movement of waterfowl, which can spread plant fragments between bodies of water. A. filiculoides will continue to invade countries where the presence of eutrophic waters, lack of natural enemies and an unregulated nursery trade will contribute to its status as a weed.
HabitatTop of page
A. filiculoides in its native areas (South America and western North America) is a plant of slow flowing streams and rivers, ponds and lakes (Reed, 1962; Lumpkin and Plucknett, 1980; Ashton, 1982). Its native range is characterised by warm, tropical climates with humid summers and mild winters. It has commonly been utilised as an ornamental in fishponds and tanks and has spread from these foci, exhibiting a weedy phenology in nutrient enriched reservoirs and roadside canals (T. Center, Senior Researcher, Aquatic Weeds, United States Department of Agriculture, personal communication).
Habitat ListTop of page
|Terrestrial||Natural / Semi-natural||Wetlands||Present, no further details||Harmful (pest or invasive)|
|Freshwater||Present, no further details||Harmful (pest or invasive)|
Hosts/Species AffectedTop of page
A. filiculoides is not generally considered a weed of crops. It is commonly grown in conjunction with rice, as A. filiculoides has the ability to fix nitrogen via an endosymbiotic blue-green alga (Wagner, 1997), acting as a green manure in rice cultivation. However, in its introduced range, A. filiculoides may impact upon trout and other fish farming (McConnachie et al., 2003). It has also been recorded to affect the growth of Potamogeton crispus L. (Janes et al., 1996).
Host Plants and Other Plants AffectedTop of page
|Potamogeton crispus (curlyleaf pondweed)||Potamogetonaceae||Other|
Growth StagesTop of page
Biology and EcologyTop of page
The chromosome number of A. filiculoides is 2n=44 (Stergianou and Fowler, 1990). This number is shared by A. pinnata, A. caroliniana, A. microphylla and A. mexicana. Hybridization has been recorded with A. microphylla under laboratory conditions (van Cat et al., 1989), the artificial hybrid having the same chromosome number as its parents (Stergianou and Fowler, 1990). Hybrids were found to produce only microsporocarps, had intermediate stem lengths, and grew better than A. microphylla in the field (van Cat et al., 1989).
Physiology and Phenology
Megaspores are produced in spring and summer, and can overwinter and survive extreme desiccation (Lumpkin and Plucknett, 1982). Megaspores germinate on the mud surface at the bottom of a waterbody, often in very shallow (0.1-0.2 m) areas. Ashton (1982) conducted detailed studies regarding A. filiculoides spore germination. The first visible sign of germination is the upward displacement of the indusium cap, due to the expansion of the first leaf. Eventually, as the second and third leaves start pushing upwards, the indusium cap is lost and the new sporophyte floats to the water surface where further development takes place. The first root appears from the base of the first leaf, and at this point the remains of the old megaspore break away from the sporophyte. Continued development results in the rapid elongation of the rhizome and the production of further leaves and roots.
A. filiculoides is able to undergo rapid vegetative reproduction throughout the year by the elongation and fragmentation of the small fronds, and under ideal conditions, the daily rate of increase can exceed 15% with the doubling time being every 4-5 days (Lumpkin and Plucknett, 1982). Under favourable environmental conditions, A. filiculoides undergoes sexual reproduction. Pairs of sporocarps are formed from a ventral lobe initial of a lateral branch. The sporocarps are of two types, male microsporocarps and female megasporocarps (Ashton, 1982; Wagner, 1997). There is usually a pair of either microsporocarps or megasporocarps, but one of each may be present (Moore, 1969). Microsporocarps are approximately 1.5 mm in diameter, containing 8-130 microsporangia (Wagner, 1997). Within each microsporangium there are 64 microspores, which are, in turn, aggregated into 3-10 massulae (Moore, 1969). Megasporocarps are approximately 0.5 mm in diameter, each producing a single megasporangium (Wagner, 1997). A single megaspore, which contains a small colony of Anabaena azollae is contained within the megasporangium (Wagner, 1997). Upon reaching maturity, both micro- and megasporocarps dehisce. Microsporangia release spongy masses of massulae into the water, which attach to megasporocarps via barbed, protruding appendages (glochidia) (Lumpkin and Plucknett, 1980). These entanglements usually sink to the bottom of a waterbody and, after a period of dormancy, the micro- and megaspores will germinate to form prothalli (Wagner, 1997). Ciliated, male gametes (antherozoids) develop in antheridia on the male thallus and female gametes (oospheres) develop in archegonia on the female thallus. After fertilization of the oospheres by the antherozoids, an embryo develops (Moore, 1969). Ashton (1982) found megaspore germination to be affected by desiccation (desiccation for >40 days greatly reduced germination); turbulence (severe turbulence lowered germination levels to a low 6%); photoperiod; light intensity; and pH (no germination occurred at pH levels <4.5 or >9.5).
A. filiculoides grows in association with the heterocystous cyanobacterium (blue-green alga) Anabaena azollae (Nostocales: Nostocaceae), within the dorsal leaf lobe cavities (Ashton and Walmsley, 1984). The alga has the ability to fix atmospheric nitrogen and is able to fulfil the nitrogen requirements of the fern making it successful in nitrogen-deficient waters (Ashton, 1982).
A. filiculoides is an obligate aquatic plant, whose growth is phosphorous limited. Climatic requirements include suitably warm months for sporocarp development, adequate radiation and light intensity for vegetative growth, and adequate amounts of rainfall to prevent its aquatic habitat from drying up. This species of tropical origin is thought to have evolved a cold-tolerant strain since its introduction into Britain (Janes, 1998b) and South Africa (McConnachie, 2003). A. filiculoides may be able to survive temperatures as low as -10ºC before death occurs.
Latitude/Altitude RangesTop of page
|Latitude North (°N)||Latitude South (°S)||Altitude Lower (m)||Altitude Upper (m)|
Air TemperatureTop of page
|Parameter||Lower limit||Upper limit|
|Absolute minimum temperature (ºC)||-10|
|Mean annual temperature (ºC)||20||28|
|Mean maximum temperature of hottest month (ºC)||18||39|
|Mean minimum temperature of coldest month (ºC)||-4||15|
RainfallTop of page
|Parameter||Lower limit||Upper limit||Description|
|Dry season duration||0||0||number of consecutive months with <40 mm rainfall|
|Mean annual rainfall||40||545||mm; lower/upper limits|
Rainfall RegimeTop of page
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Pseudolampsis darwinii||Herbivore||Plants|Leaves||to genus||Hill (1999)|
|Pseudolampsis guttata||Herbivore||Plants|Leaves||Hill and Oberholzer (2002)|
|Stenopelmus brunneus||Herbivore||Plants|Leaves||to species|
|Stenopelmus rufinasus||Herbivore||Plants|Leaves||to genus||Hill (1998)||South Africa|
Notes on Natural EnemiesTop of page
Host records from around the globe show that the genus Azolla is attacked by generalist herbivores and that very few specialist insect species have evolved on these plants (Hill, 1997). However, four beetle species, the weevils Stenopelmus rufinasus and S. brunneus and the two flea beetles Pseudolampsis guttata and P. darwinii, appear to have specialized on the genus Azolla (Richerson and Grigarick, 1967; Habeck, 1979; Hill, 1999) and were identified as potential biological control agents for A. filiculoides in South Africa (Hill, 1997). Following host range testing, Stenopelmus rufinasus was released in 1997 as a biocontrol of A. filiculoides in South Africa (McConnachie et al., 2004).
Means of Movement and DispersalTop of page
A. filiculoides propagates both asexually and sexually. Both spores and plant fragments are dispersed long distances along river systems.
Waterfowl, amphibians and rodents are thought to spread small fragments of the plant that adhere to their bodies.
A. filiculoides has been introduced in tropical regions worldwide as a green manure in rice cultivation.
A. filiculoides was intentionally introduced into South Africa as a fish pond plant in 1948 (Jacot-Guillarmod, 1979) and by the early 1980s it was thought to have invaded every river system in South Africa. The plant was also deliberately introduced into south-east UK at the end of the nineteenth century as an ornamental plant (Janes, 1998a) and is now naturalised in numerous still and slow-flowing waters (Preston and Croft, 1997). There are also instances of research collections that have escaped from culture; A. filiculoides has been spread around the world as a model plant to study the Azolla-Anabaena symbiosis (Carrapiço, 2010)
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Growing medium accompanying plants||weeds/seeds||Yes|
|Plant parts not known to carry the pest in trade/transport|
|Fruits (inc. pods)|
|Stems (above ground)/Shoots/Trunks/Branches|
|True seeds (inc. grain)|
Impact SummaryTop of page
|Fisheries / aquaculture||Negative|
Economic ImpactTop of page
The economic impact of A. filiculoides in South Africa was examined by McConnachie et al. (2003). Thick mats on reservoirs and slow-moving waterbodies caused economic losses to water-users. Among those water-uses most seriously affected were farming (71%), recreational (24%), and municipal (5%). On average, A. filiculoides was found to cause on-site damages of US$589 per hectare per year.
Environmental ImpactTop of page
In eutrophic water systems, A. filiculoides grows rapidly, easily outcompeting indigenous vegetation. Decaying root and leaf matter below a mat of A. filiculoides, and the lack of light penetration, creates an anaerobic environment. Not only can very little survive under such conditions, but the quality of drinking water is reduced, caused by bad odours, colour and turbidity (Hill, 1997). Cases have been reported where both livestock and game farmers have lost animals due to them refusing to drink from infested waterbodies or drowning as a result of mistaking the mat for solid ground. The weed also reportedly increases water loss through evapotranspiration and promotes the development of waterborne, water-based and water-related diseases (Hill, 1997).
Impact on Biodiversity
A. filiculoides infestations may form thick mats (5-20 cm thick), on waterbodies up to 10 hectares in size (McConnachie et al., 2003). Such infestations have been shown to severely impact the biodiversity of aquatic ecosystems and have serious implications for all aspects of water utilisation (Gratwicke and Marshall, 2001).
One of the last remaining habitats of the endangered fish species, the eastern Cape rocky (Sandelia bainsii Castelnau, 1861; Anabantidae) in South Africa, had become so overgrown with the weed that had the biological control programme not been so successful, S. bainsii faced extinction.
Threatened SpeciesTop of page
Social ImpactTop of page
Primarily, social impacts of A. filiculoides have centred around the reduction of useful water surface area for recreation (fishing, swimming and water skiing) and water transport.
Risk and Impact FactorsTop of page
- 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 human health
- Negatively impacts animal health
- Negatively impacts tourism
- Reduced amenity values
- Reduced native biodiversity
- Highly likely to be transported internationally accidentally
- Highly likely to be transported internationally deliberately
- Difficult to identify/detect as a commodity contaminant
- Difficult to identify/detect in the field
UsesTop of page
Members of the genus Azolla are utilized throughout the world for a wide variety of purposes besides its widespread uses as an ornamental in fish ponds and tanks (Lumpkin and Plucknett, 1980; 1982). A. filiculoides is used as a green manure in rice paddies, mainly in Asia, as an inhibitor of weed growth in rice cultivation in China and Vietnam (Kröck and Alkämper, 1991), and as an alternative high protein fodder for cattle, swine, poultry and fish, and possibly as an alternative food source for humans, again, mainly in Asia. It has also been used as a nitrate-rich compost which potentially increases soil organic nitrogen levels and cation exchange capacity. It is used for purification of water, removal of heavy metals (Sanyahumbi et al., 1998) and removal of nitrogen and phoshorous from wastewater (Forni et al., 2001). It has also been used variously as an ingredient in soap production, a cure for sore throats and as a control for mosquitoes in southern India as complete mats disrupt larval development (Rajendran and Reuben, 1991).
Similarities to Other Species/ConditionsTop of page
A. filiculoides is difficult to distinguish from A. microphylla, A. mexicana, A. caroliniana and A. cristata due to the variable morphology of the plant modified by interacting environmental influences (Ashton, 1982). Cytological studies are the most reliable method of separating these species (Stergianou and Fowler, 1990; Saunders and Fowler, 1992; 1993). Under field conditions, however, the most reliable measure of separating these species would be by means of their sporocarp structure (Svenson, 1944; Sweet and Hills, 1971). A. filiculoides: glochidia not septate, or rarely with 1 or 2 septae at the apex, microsporangia 35-100 in an indusium, massulae 4-6, megasporangia with raised, irregular hexagonal markings; A. caroliniana: glochidia not septate, microsporangia 8-40 in an indusium; A. mexicana: glochidia multi-septate, microsporangia usually with 4 massulae, megaspore pitted; A. microphylla: glochidia multi-septate, megaspore smooth. For comparisons including A. cristata, see Evrard and Van Hove (2004).
Prevention and ControlTop 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.
Even though A. filiculoides has been used as fodder for pigs, poultry and cattle (Lumpkin and Plucknett, 1982), control through grazing may not be achieved because of its aquatic habit. However, A. filiculoides may actually be a preferred forage for herbivorous fish, even over most other aquatic weeds (Edwards, 1974).
Small infestations of the weed in accessible areas may be removed using rakes and fine meshed nets (Hill, 1997). The disadvantage of mechanical control, however, is that under ideal conditions, the weed can double itself every 4-5 days (Lumpkin and Plucknett, 1982). Hence a concerted effort would be required just to keep up with the daily production of even a small infestation, and even if eradication were achieved, re-establishment from spores resident in the waterbody substrate would be inevitable. Ashton (1992) noted that A. filiculoides was susceptible to fragmentation from physical disturbance, and that the fragments were highly sensitive to high light intensity and were killed by direct sunlight. He thus suggested the use of a mechanical agitator or stirrer to provide enough turbulence to break up the plants. The cost, however, of such an approach (even on a small scale) would be prohibitive.
Chemicals proposed to control A. filiculoides include glyphosate (Steyn et al., 1979; Ashton, 1992), paraquat and diquat (Axelsen and Julien, 1988), and kerosene mixed with a surfactant (Diatloff and Lee, 1979). However, paraquat is now banned in the EU, Switzerland and a number of other countries. Diquat use in the EU is restricted to terrestrial treatments and can no longer be authorised for aquatic weed control. Glyphosate is toxic to fish and algae and the water cannot be used for irrigation or stock until the herbicide has broken down.
South Africa is the only country that has initiated a classical biological control programme against A. filiculoides. Four insect species were identified as potential biological candidates - all frond-feeding beetles: Pseudolampsis guttata Leconte (Chrysomelidae), P. darwinii Sherer, Stenopelmus rufinasus Gyllenhal (Curculionidae) and S. brunneus Hustache . All species do extensive damage to the plants in the country of origin (Hill, 1997). The biology and host range of P. guttata was investigated by Hill (2002), and although it was found to be fairly damaging, the weevil S. rufinasus was identified as the most suitable of the four for release in South Africa, and was imported into quarantine for host-specificity screening. The weevil was released in 1997 and results have been dramatic, causing local extinction of A. filiculoides at the sites where it was released (McConnachie et al., 2004). The surface area of weed controlled totalled 203.5 ha and infested sites were controlled in approximately seven months on average (in a range of 3-11 months). Five years after the release of the weevil, A. filiculoides no longer poses a threat to aquatic ecosystems in South Africa and its effects on the utilization of water resources have been significantly reduced (McConnachie et al., 2003). S. rufinasus has been released in Mozambique and Zimbabwe with material provided by South Africa, has established, and is proving a successful biocontrol agent in these countries also (Cilliers et al., 2003).
S. rufinasus has been present in the UK since it was first reported there by Janson (1921) and was probably brought into Europe with the plant. Since then it has been recorded in Ireland, France, Belgium, the Netherlands and Spain (Pratt et al., 2013). This agent has been used as in an augmentive manner in England and Ireland and is under consideration for mass-rearing and releasing in the Netherlands, France and Belgium (Pratt et al., 2013).
Because of the success of the biological control of A. filiculoides, there has been no need to develop an integrated approach.
Case Study: A. filiculoides in South Africa
A. filiculoides was first recorded in South Africa in 1948 from the Oorlogspoort River in the Northern Cape Province of South Africa (30°37’58.22”S 25°21’28.89”E), and by 1999 it was recorded from 152 sites in South Africa, largely in the Free State Province.
Owing to the adverse environmental and economic effects of the weed, a biological control programme was initiated with the importation of the weevil Stenopelmus rufinasus Gyllenhal (Coleoptera: Curculionidae) from Florida, USA, in 1995. Following host-specificity testing, the weevils were released in South Africa in 1997. By 2004, nearly 25,000 weevils had been released throughout South Africa, and their feeding damage has resulted in local extinctions of A. filiculoides from the majority of sites that were surveyed at the time. It took, on average, ten months for a site to be cleared after release of the weevils. The dispersal abilities of the weevils were originally underestimated, but they are capable of dispersing up to 350 km unaided.
Just five years after the release of the weevils, A. filiculoides was no longer considered a threat to South African waterbodies (McConnachie et al., 2004). These successes were carefully monitored between 1999 and 2006, documenting the rapid control of the plant, and proving that just four years after the studies by McConnachie et al. (2004) the weevils had succeeded in controlling A. filiculoides at every site where they had been released (Hill et al., 2008).
Country-wide surveys from 2008 produced further evidence of the success of this program. In 2010, of the 102 A. filiculoides sites investigated (40% of water weed sites surveyed), the weed was present at 19 (19%) of these sites, and S. rufinasus was recorded from 14 (73%) of the infested sites. A. filiculoides is no longer a significant problem in South Africa, and where it does occur, S. rufinasus is usually present. Biological control of A. filiculoides is now widely regarded as the most successful biological control programme against an invasive alien weed in South Africa (Coetzee et al., 2011).
Field surveys indicated that the agent had started to use another host thought to be the native A. pinnata subsp. africana, however a recent study by Madeira et al. (2016) showed that it was in fact feeding on A. cristata, another introduced fern that is more closely related to A. filiculoides.
Gaps in Knowledge/Research NeedsTop of page
Further research on the taxonomy of the genus Azolla is on-going (Madeira et al., 2013, 2016).
ReferencesTop of page
Ashton PJ, 1982. The autecolgy of Azolla filiculoides Lamarck with special reference to its occurrence in the Hendrik Verword Dam catchment area. PhD thesis, Rhodes University, South Africa
Ashton PJ, 1992. Azolla infestations in South Africa: history of the introduction, scope of the problem and prospects for management. Water Quality Information Sheet
Ashton PJ, Walmsley RD, 1984. The taxonomy and distribution of Azolla species in southern Africa. Botanical Journal of the Linnean Society, 89:239-247
Bailey FM, 1902. The Queensland flora, Pt VI. Brisbane, Australia: HJ Didams and Co
Brunker JP, 1949. Azolla filiculoides Lam. In Co. Wicklow. Irish Naturalists' Journal, 9:340
Carrapico F, 2010. Azolla as a Superorganism. Its Implication in Symbiotic Studies. Symbioses and Stress Cellular Origin, Life in Extreme Habitats and Astrobiology, 17:225-241
Chevalier A, 1926. La culture des Azolla pour la nourriture des animaux de basse-cour et comme engrais vert pour les riziers. Rev. Bot. Appl. Agr. Trop., 6:356-360
Chikwenhere GP, 2001. Current strategies for the management of water hyacinth on the Manyame River System in Zimbabwe. Biological and integrated control of water hyacinth: Eichhornia crassipes. Proceedings of the Second Meeting of the Global Working Group for the Biological and Integrated Control of Water Hyacinth, Beijing, China, 9-12 October 2000, 105-108
Cilliers CJ, Hill MP, Ogwang JA, Ajuonu O, 2003. Aquatic weeds in Africa and their control. In: Neuenschwander P, Borgemeister C, Langewald J, eds. Biological Control in IPM Systems in Africa. Wallingford, UK: CAB International, 161-178
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Langa SDF, 2013. PhD Thesis., Grahamstown, South Africa: Rhodes University.
Large MF, Braggins JE, 1993. Spore morphology of the New Zealand Azolla filiculoides Lam. In: New Zealand Journal of Botany, 31 419-423.
Lawalree A, 1964. Azollaceae. In: Flora Europaea, [ed. by Tutin TG, Haywood VH, Vallentyne DH, Walters SM, Webb DA]. UK: Cambridge University Press.
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Magda S A, Ghazi I M, Atalla K M, Nahed M A, 2002. Influence of salinity and source of water on growth and protein content of Azolla pinnata and Azolla filiculoides under Egyptian conditions. Annals of Agricultural Science, Moshtohor. 40 (2), 871-884.
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Seto K, Nasu T, 1975. Discovery of fossil Azolla massulae from Japan and some notes on recent Japanese species [In Japanese]. In: Bull. Osaka Mus. Nat. Hist. 29 51-60.
Sourek J, 1958. (Azolla filiculoides Lam. - neue eingeschleppte Art in der CSR). In: Preslia, 30 84-85.
Twyman ES, Ashton PJ, 1972. An autecological investigation of the aquatic fern (Azolla species) in relation to the Hendrik Verwoed Dam. In: The Civil Engineer in South Africa, 14 88-89.
USDA-ARS, 2003. Hedychium flavescens. In: Germplasm Resources Information Network (GRIN). Online Database, Beltsville, USA: National Germplasm Resources Laboratory. http://www.ars-grin.gov/cgi-bin/npgs/html/tax_search.pl
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Witt A, Luke Q, 2017. Guide to the naturalized and invasive plants of Eastern Africa. [ed. by Witt A, Luke Q]. Wallingford, UK: CABI. vi + 601 pp. http://www.cabi.org/cabebooks/ebook/20173158959 DOI:10.1079/9781786392145.0000
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27/06/2014 Datasheet updated by:
Martin Hill, Rhodes University, South Africa
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