Invasive Species Compendium

Detailed coverage of invasive species threatening livelihoods and the environment worldwide

Datasheet

Asparagus asparagoides
(bridal creeper)

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Datasheet

Asparagus asparagoides (bridal creeper)

Pictures

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PictureTitleCaptionCopyright
Asparagus asparagoides; foliage and below-ground biomass, consisting of fleshy tubers attached to a network of rhizomes.
TitleFoliage and below-ground biomass
CaptionAsparagus asparagoides; foliage and below-ground biomass, consisting of fleshy tubers attached to a network of rhizomes.
CopyrightCSIRO
Asparagus asparagoides; foliage and below-ground biomass, consisting of fleshy tubers attached to a network of rhizomes.
Foliage and below-ground biomassAsparagus asparagoides; foliage and below-ground biomass, consisting of fleshy tubers attached to a network of rhizomes.CSIRO

Identity

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Preferred Scientific Name

  • Asparagus asparagoides (L.) Druce, 1914

Preferred Common Name

  • bridal creeper

Other Scientific Names

  • Asparagus medeoloides (L.f.) Thunb. (1794)
  • Asparagus medioloides (L. fil.) Thunb.
  • Dracaena medeoloides L. f. (1782)
  • Elachanthera sewelliae F. Muell.
  • Luzuriaga sewelliae (F. Muell.) K. Krause (1930)
  • Medeola asparagoides L. (1753)
  • Myrsiphyllum asparagoides (L.) Willd. (1808)

International Common Names

  • English: African asparagus fern; bridal creeper; bridal veil creeper; cape smilax; florist's smilax; smilax; smilax asparagus; smilax asparagus
  • French: liane asperge

Local Common Names

  • Australia: bridal veil creeper
  • New Zealand: smilax
  • South Africa: gnarboola; krulkransie; narbas
  • USA: African asparagus fern

EPPO code

  • ASPAS (Asparagus asparagoides)

Summary of Invasiveness

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The South African native plant A. asparagoides is a dense, scrambling vine capable of invading a range of habitats in warm temperate climates (Morin et al., 2006a). It was introduced to Australia as a garden ornamental in the mid-1800s and within a century had extensively naturalized. It was ranked in the top 20 Weeds of National Significance in Australia in the late 1990s (Thorp and Lynch, 2000) and is also reported as invasive in New Zealand (Roy et al., 2004). It invades both disturbed and undisturbed natural ecosystems, where it quickly dominates and smothers understorey vegetation and changes the structure, floristic composition and ecology of the system (Turner et al., 2008b). It is not known to invade agricultural systems, except for citrus orchards in irrigation areas of Australia, where it smothers trees and displaces citrus roots leading to reduced fruit production (Kwong and Holland-Clift, 2004).

Taxonomic Tree

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  • Domain: Eukaryota
  •     Kingdom: Plantae
  •         Phylum: Spermatophyta
  •             Subphylum: Angiospermae
  •                 Class: Monocotyledonae
  •                     Order: Liliales
  •                         Family: Liliaceae
  •                             Genus: Asparagus
  •                                 Species: Asparagus asparagoides

Notes on Taxonomy and Nomenclature

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In their latest Australian treatment of the Asparagaceae, Clifford and Conran (1987) included Asparagus asparagoides in the genus Myrsiphyllum, based on Obermeyer’s (1984) revision of the family, which divided it into three genera, Asparagus, Protasparagus and Myrsiphyllum. In a subsequent revision, Malcomber and Demissew (1993) concluded that the Asparagaceae contains only one genus, Asparagus, with two subgenera, Asparagus (which includes Obermeyer’s Protasparagus) and Myrsiphyllum. The decision that the family should comprise a single genus was upheld by Fellingham and Meyer (1995), but they argued that the subgeneric status of Asparagus and Myrsiphyllum is not warranted. A recent molecular phylogeny of Asparagus, however, found evidence that southern African species are potentially paraphyletic, supporting sub-division of Asparagus into more than three groups (Fukuda et al., 2005).

Description

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The following description was compiled mostly from Obermeyer (1984), Clifford and Conran (1987), Scott and Kleinjan (1991), Kleinjan and Edwards (1999) and Parsons and Cuthbertson (2001).

A. asparagoides is a geophyte with extensive, perennial, below-ground storage organs and above-ground parts that die back annually or when conditions are unfavourable. It produces thin, wiry, twisting shoots, slightly woody at the base and up to 6 m long when offered support. Shoots emerging from the below-ground root system entwine with each other and surrounding vegetation, allowing them to climb understorey shrubs and small trees.

It produces leaf-like stems called cladodes or phylloclades, arising in the axils of reduced, scale-like leaves. The cladodes are stalkless, broadly ovate to lanceolate but sharply pointed with a smooth or minutely denticulate margin, 10–70 mm long, 4–30 mm wide, dark glossy-green when growing in shade, but dull and light green in exposed locations and have a delicate parallel venation with no prominent mid-vein. They are solitary and alternate along the stem, or borne in groups on short side branches.

Flowers are sweetly-scented, 8–9 mm wide and 5–6 mm long when fully expanded. They are borne on 3–8 mm long and slightly bent pedicels, singly or in pairs in the axils of the reduced scale-like leaves. The six tepals (term used when petals and sepals are similar in appearance) are greenish-white, turned backward and fused into a tube in the lower half. The six stamens are connivent (touching but not fused) and slightly shorter than the tepals.

Fruits are globular berries, 6–10 mm wide, initially green and ripening red. They generally contain 0–4 black, shiny, spherical or ovoid seeds.

The below-ground biomass consists of a cylindrical, slender (about 5 mm wide), branching rhizome (underground stem with shoot buds) growing parallel to the soil surface and bearing numerous, radially-arranged, spindle-shaped, fleshy tubers, 25–42 mm long and 8–20 mm wide, often continuing as roots. Rhizomes and tubers of large plants entwine, forming a dense mat about 5–10 cm thick just below the soil surface.

Plant Type

Top of page Herbaceous
Perennial
Seed propagated
Vegetatively propagated
Vine / climber

Distribution

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The native range of A. asparagoides is southern Africa, but it is also recorded from Namibia and further north in tropical Africa (Obermeyer, 1984). Kleinjan and Edwards (1999), however, pointed out that the veracity of these northern herbarium records cannot be ascertained due to the absence of tubers on specimens. A. asparagoides is widely distributed in South Africa and occurs in the three main rainfall zones (winter, summer and even rainfall) (Kleinjan and Edwards, 1999). It does not occur in the dry interior of the Western Cape and Northern Cape Provinces or the hot subtropical coast of KwaZulu-Natal.

A. asparagoides is naturalised in all states of Australia, but not in the Northern and Australian Capital Territories (Morin et al., 2006a). It is most prevalent and invasive in the temperate and Mediterranean regions of southern Australia. Climate modelling to predict its potential distribution in Australia indicated that slight northern expansions could be expected along the east and west coasts of the mainland and on the north and east coasts of Tasmania (Scott and Batchelor, 2006).
 
A. asparagoides is also considered an invasive plant in New Zealand (Roy et al., 2004) where it is most common in the northern regions of Northland, Auckland, Waikato, Bay of Plenty and Gisborne (Harman et al., 2008). It is naturalised in some counties of California, USA, and although infestations are not large at this stage, managers have been alerted to its potential invasiveness (Calflora, 2010). Climate modelling showed that large areas of the Californian coast are suitable for A. asparagoides establishment (Randall and Lloyd, 2002). A. asparagoides is recorded as locally naturalised but not yet invasive in several islands around the world including Maui (Hawaii), Sicily, Malta and Corsica (Valdés, 1980; Wagner et al., 2005; Global Compendium of Weeds, 2007; Royal Botanic Garden Edinburgh, 2010). It is recorded from Guatemala, Mexico and Slovenia, but limited information is available (Global Compendium of Weeds, 2007; Missouri Botanical Garden, 2008).

 

Distribution Table

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

Continent/Country/RegionDistributionLast ReportedOriginFirst ReportedInvasiveReferenceNotes

Africa

EthiopiaAbsent, unreliable recordKleinjan and Edwards, 1999; USDA-ARS, 2010; EPPO, 2014The veracity of records from tropical Africa cannot be ascertained due to the absence of tubers on herbarium specimens
KenyaPresentEPPO, 2014
LesothoWidespreadNative Not invasive Kleinjan and Edwards, 1999; EPPO, 2014
MalawiPresentEPPO, 2014
MoroccoPresentEPPO, 2014
NamibiaAbsent, unreliable recordObermeyer, 1984; Kleinjan and Edwards, 1999; EPPO, 2014The veracity of records from tropical Africa cannot be ascertained due to the absence of tubers on herbarium specimens
South AfricaWidespreadNative Not invasive Kleinjan and Edwards, 1999; EPPO, 2014
Spain
-Canary IslandsPresentEPPO, 2014
SwazilandWidespreadNative Not invasive Kleinjan and Edwards, 1999; EPPO, 2014
TanzaniaPresentEPPO, 2014
TunisiaPresentEPPO, 2014
UgandaPresentEPPO, 2014
ZambiaPresentEPPO, 2014
ZimbabwePresentEPPO, 2014

North America

MexicoPresentIntroducedMissouri Botanical Garden, 2010; EPPO, 2014
USARestricted distributionEPPO, 2014
-CaliforniaLocalisedIntroducedCalflora, 2010; EPPO, 2014
-HawaiiPresent, few occurrencesIntroducedWagner et al., 2005; EPPO, 2014Reported from East Maui

Central America and Caribbean

GuatemalaPresentIntroducedMissouri Botanical Garden, 2010; EPPO, 2014

South America

ArgentinaPresentEPPO, 2014
UruguayPresentEPPO, 2014

Europe

FrancePresentIntroduced Not invasive Global Compendium of Weeds, 2007; EPPO, 2014Corsica
-CorsicaPresentEPPO, 2014
ItalyPresentIntroduced Not invasive Royal Botanic Garden Edinburgh, 2010; EPPO, 2014Sicily
-SardiniaAbsent, no pest recordEPPO, 2014
-SicilyPresentEPPO, 2014
MaltaPresentIntroduced Not invasive Royal Botanic Garden Edinburgh, 2010; EPPO, 2014
PortugalPresentIntroduced Not invasive Royal Botanic Garden Edinburgh, 2010; EPPO, 2014Including Azores Archipelago
-AzoresPresentEPPO, 2014
-MadeiraPresentEPPO, 2014
SloveniaAbsent, invalid recordGlobal Compendium of Weeds, 2007; EPPO, 2014
SpainPresentIntroduced Not invasive Valdés, 1980; EPPO, 2014Canary Islands

Oceania

AustraliaLocalisedIntroducedMorin et al., 2006a; EPPO, 2014Including Lord Howe Island
-New South WalesLocalisedIntroduced Invasive Morin et al., 2006a; EPPO, 2014
-QueenslandPresent, few occurrencesIntroduced Invasive Morin et al., 2006a; EPPO, 2014
-South AustraliaLocalisedIntroduced Invasive Morin et al., 2006a; EPPO, 2014
-TasmaniaLocalisedIntroduced Invasive Morin et al., 2006a; EPPO, 2014
-VictoriaLocalisedIntroduced Invasive Morin et al., 2006a; EPPO, 2014
-Western AustraliaLocalisedIntroduced Invasive Morin et al., 2006a; EPPO, 2014
New ZealandLocalisedIntroduced Invasive Roy et al., 2004; EPPO, 2014

History of Introduction and Spread

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Over the past two centuries, A. asparagoides has been deliberately introduced as a garden ornamental in many temperate regions of the world. The 1857 plant catalogue of William MacArthur represents the earliest known record of A. asparagoides being available for sale in Australia (Mulvaney, 1991). A. asparagoides was also listed in other nursery catalogues in the 1870s and 1880s (Mulvaney, 1991; Brookes and Barley, 1992) and recorded as a plant growing in the Adelaide and Melbourne Botanic Gardens in 1871 and 1883, respectively (Scott, 1995).

By the early 1900s, A. asparagoides was widely used by florists for decoration and as greenery in bridal bouquets (Scott, 1995). It was also extensively used as a garden ornamental at that time. A. asparagoides’ popularity with florists and gardeners was short-lived and declined steadily during the first part of the twentieth century. It was first recorded as naturalised in Victoria in 1886 (Parsons and Cuthbertson, 2001), although the first herbarium specimen of naturalised plants for that State was collected in 1943 (Clifford and Conran, 1987). The earliest herbarium records of naturalised plants for South Australia and Western Australia were made in 1937 and 1960, respectively (Scott, 1995). Its initial distribution in the wild was clearly linked to locations of early settlements (Stansbury and Scott, 1999).

A. asparagoides naturalized in New Zealand between 1900 and 1940 (Esler and Astridge, 1987).

Introductions

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Introduced toIntroduced fromYearReasonIntroduced byEstablished in wild throughReferencesNotes
Natural reproductionContinuous restocking
Australia South Africa 1850s Nursery trade (pathway cause) Yes Mulvaney (1991) Possibly introduced via Europe

Risk of Introduction

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The risk of introduction of new genotypes of A. asparagoides to Australia, which could jeopardize the effectiveness of biological control, led to its listing as a prohibited species under Australian plant quarantine legislation (ICON, 2010). Its entry is also prohibited in New Zealand (Plants Biosecurity Index, 2010). Risks of deliberate introduction in other countries are high due to A. asparagoides’ ornamental attributes, but naturalization is unlikely to occur outside of temperate regions.  

Trade, movement and sale of A. asparagoides is prohibited in all states and territories in Australia (Morin et al., 2006a).

Habitat

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In South Africa, A. asparagoides mainly occurs as a minor understorey species (Kleinjan and Edwards, 1999). In contrast, it is capable of invading a variety of habitats in warm temperate climates of Australia and New Zealand including coastal heath on sandy dunes, woodlands or forests, creek and river banks, swamps, dry coastal vegetation, mallee shrubland, dry and damp sclerophyll open-forest, and littoral rainforest (Roy et al., 2004; Morin et al., 2006a). It prefers shaded or part-shaded habitat, but can also grow in hind dunes on exposed beaches, coastal cliffs and amongst shrubs closer to shoreline in sheltered bays. Invasion of new sites appears unrelated to disturbance events (Raymond, 1999; Siderov et al., 2006). Stansbury and Scott (1999), however, found via a questionnaire distributed to landholders in Western Australia that there was a weak association between properties that had a long history of disturbance through clearing of native vegetation and the presence of A. asparagoides.

Habitat List

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CategorySub-CategoryHabitatPresenceStatus
Terrestrial
 
Terrestrial – ManagedManaged forests, plantations and orchards Principal habitat Harmful (pest or invasive)
Managed forests, plantations and orchards Principal habitat Natural
Disturbed areas Principal habitat Harmful (pest or invasive)
Disturbed areas Principal habitat Natural
Rail / roadsides Principal habitat Harmful (pest or invasive)
Rail / roadsides Principal habitat Natural
Urban / peri-urban areas Secondary/tolerated habitat Natural
Urban / peri-urban areas Secondary/tolerated habitat Productive/non-natural
Terrestrial ‑ Natural / Semi-naturalNatural forests Principal habitat Harmful (pest or invasive)
Natural forests Principal habitat Natural
Riverbanks Principal habitat Harmful (pest or invasive)
Riverbanks Principal habitat Natural
Scrub / shrublands Principal habitat Harmful (pest or invasive)
Scrub / shrublands Principal habitat Natural
Littoral
Coastal areas Principal habitat Harmful (pest or invasive)
Coastal dunes Secondary/tolerated habitat Natural

Hosts/Species Affected

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A. asparagoides has been observed to eliminate or restrict growth of a wide range of native plant species (see Impact: Environmental).

It does not invade pastures as it cannot withstand constant grazing. It is also not an invader in broad-acre cropping situations probably because of the limited seed input in such environments, preference for shaded habitats and the vulnerability of seedlings to cultivation. It invades pine plantations, but it is not perceived to have a significant impact on tree growth. It is, however, troublesome in citrus orchards in Australia, where it smothers trees, displaces citrus roots and reduces fruit production (Kwong and Holland-Clift, 2004).

 

Host Plants and Other Plants Affected

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Plant nameFamilyContext
CitrusRutaceaeMain

Growth Stages

Top of page Flowering stage, Fruiting stage, Vegetative growing stage

Biology and Ecology

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Genetics
 
Comparison of sequences of non-coding DNA regions from the chloroplast (trnL-trnF) and nuclear ribosomal (ITS1 and ITS2) genomes of South African and Australian accessions of A. asparagoides, identified the Western Cape Province of South Africa as the likely geographical source of founder populations in Australia (L Morin, CSIRO Entomology, Australia, personal communication, 2010).

Chromosome number 2n = 20.
 
Reproductive Biology
 
A. asparagoides’ flowers are bisexual and self-compatible. Cross-pollination has been demonstrated in hand-pollinated flowers, but the level of outcrossing versus selfing still remains to be determined (Raymond, 1999). Honeybees have been observed visiting flowers in large numbers, collecting both pollen and nectar (Raymond, 1999).
 
Shoots that emerge from well-established plants early in the growing season are the only ones that will produce floral structures and fruits, providing they can climb on a support (Raymond, 1999; Stansbury et al., 2007). Buds flower and wither within two weeks and, by the third week swelling of the ovary is usually visible. Fruit set is usually low < 50%) probably due to a combination of climatic or site-specific factors such as high intraspecific competition and stress among flowering shoots, extreme shading, herbivory on buds and flowers and mouldiness of developing fruits leading to abortion (Raymond, 1999).
 
Physiology and Phenology
 
A. asparagoides seeds have an after-ripening requirement that ensures they do not germinate too soon after they are produced. Passage of seeds through the gut of silvereye (Zosterops lateralis) birds was observed to reduce the after-ripening period and enhance germination compared to un-passed seeds (Raymond, 1999). Mature seeds can germinate over a wide range of temperatures (10–30°C) in both constant light or darkness, but germination is optimal at lower temperatures of approximately 15–20°C (Raymond, 1999). Willis et al. (2003), however, observed a strong inhibitory effect of light on germination when fluctuating light and temperature regimes were used. The effect of smoke and heat shock treatment (90°C for 10 min) on germination of seeds has been investigated, but results of different trials have been contradictory, highlighting the need for further work in this area (Willis et al., 2003).
 
Following rapid growth and establishment of shoots at the beginning of the growing season, A. asparagoides diverts some of its photosynthates towards tuber production. The above-ground biomass is positively correlated to its below-ground biomass (Kleinjan et al., 2004a). Once plants establish extensive below-ground reserves they can produce shoots at the onset of cooler temperatures, even in the absence of available external moisture (Kleinjan and Edwards, 1999; Raymond, 1999).
 
In South Africa, A. asparagoides adapts its phenology to different seasonal rainfall patterns (Kleinjan and Edwards, 1999). It behaves similarly in Australia, where its major distribution is in winter-rainfall dominant regions. In most areas, shoots emerge from the below-ground rhizome either slightly before or with the onset of the first late-summer or autumn rains. Seeds readily germinate throughout autumn and winter if conditions and microhabitat are adequate. Seedlings have a good chance of surviving over the dry summer period providing they have had time to develop some below-ground reserves (Raymond, 1999).
 
In Australia, flowering generally occurs in late winter to early spring (Raymond, 1999), while in South Africa flowers are produced as early as July (Kleinjan and Edwards, 1999). Fruits develop in the spring, mature into red berries in late spring to late summer, depending on the region, and can be retained on senescing plants for several months. Above-ground biomass begins to senesce in mid to late spring. The onset of senescence is earlier when plants are exposed to more direct sunlight. Shoots typically senesce from the apex down and take several weeks to die back completely. The below-ground biomass enables plants to survive until the next growing season.
 
Associations
 
Greater accumulation of litter was observed in habitats invaded by A. asparagoides compared to non-invaded native areas in South Australia (Stephens, 2005). This difference was probably due to litter becoming trapped in the climbing stems and foliage. Despite the significant changes in habitat and decrease in plant diversity due to invasion by A. asparagoides, Stephens (2005) found a very abundant and diverse arthropod and wasp community in both native and invaded habitats. Indeed, a higher number of soil/litter-associated pollinators of the endangered orchid, Pterostylis bryophila, were recorded at the invaded sites (Stephens et al., 2003). It appeared that A. asparagoides provided a favorable habitat for these specific pollinators and did not affect their movement or pollination efficiency within the invaded habitat. These study sites, however, were not completely colonized by A. asparagoides and consequently the plant may not yet have caused a habitat change significant enough to detect flow-on negative effects on the arthropod community.
 
Regeneration of native communities after removal of A. asparagoides may be slow where few propagules of native plants are left in the soil. Below-ground biomass may also impede seedling establishment of native species by occupying all of the space available in the substrate or by changing soil chemistry. Turner and Virtue (2006) observed that dead rhizomes and tubers were still present eight years after A. asparagoides had been killed. In another study performed in Western Australia, the initial decomposition of underground biomass was rapid with about 40% loss of dry weight occurring during the first three months after burial (Turner et al., 2006). Subsequent decomposition slowed dramatically with no significant change for the next six months.
 
Environmental Requirements

A. asparagoides is particularly vigorous in soils with high moisture content and areas high in nutrients. It grows best at sites with high levels of available nitrates, potassium and iron (Stansbury, 1999). A recent study in Western Australia showed that soils where A. asparagoides dominates have higher levels of available phosphorus than nearby non-invaded areas (Turner, 2008). It is unknown at this stage whether A. asparagoides originally invaded phosphorus-rich areas or if it is modifying the soil environment.

A. asparagoides can grow in areas where rainfall is sub-optimal providing it is irrigated (e.g. private gardens or citrus orchards), or in close proximity to watercourses or in moist gullies (Stansbury and Scott, 1999).

Climate

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ClimateStatusDescriptionRemark
Cf - Warm temperate climate, wet all year Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year
Cs - Warm temperate climate with dry summer Preferred Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers
Cw - Warm temperate climate with dry winter Preferred Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)

Latitude/Altitude Ranges

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Latitude North (°N)Latitude South (°S)Altitude Lower (m)Altitude Upper (m)
42 42

Air Temperature

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Parameter Lower limit Upper limit
Absolute minimum temperature (ºC) -5
Mean annual temperature (ºC) 11 18
Mean maximum temperature of hottest month (ºC) 20 33
Mean minimum temperature of coldest month (ºC) -2 9

Rainfall

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ParameterLower limitUpper limitDescription
Dry season duration06number of consecutive months with <40 mm rainfall
Mean annual rainfall2602100mm; lower/upper limits

Rainfall Regime

Top of page Summer
Winter

Soil Tolerances

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Soil drainage

  • free

Soil reaction

  • acid
  • alkaline
  • neutral

Soil texture

  • light
  • medium

Special soil tolerances

  • infertile
  • saline
  • shallow

Natural enemies

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Natural enemyTypeLife stagesSpecificityReferencesBiological control inBiological control on
Agrotis Herbivore Leaves not specific Kleinjan and Edwards, 2006
Brachycerus Herbivore Leaves not specific Kleinjan and Edwards, 2006
Crioceris Herbivore Leaves to species Witt and Edwards, 2002
Cucullia terensis Herbivore Leaves not specific Kleinjan and Edwards, 2006
Euplexia augens Herbivore Leaves not specific Kleinjan and Edwards, 2006
Eurytoma Herbivore Seeds to genus Kleinjan and Edwards, 2006
Hespera Herbivore Leaves not specific Kleinjan and Edwards, 2006
Lycophotia Herbivore Leaves not specific Kleinjan and Edwards, 2006
Puccinia myrsiphylli Pathogen Leaves to species Kleinjan et al., 2004b
Zalaca snelleni Herbivore Fruits/pods not specific Kleinjan and Edwards, 2006
Zygina Herbivore Leaves to species Morin and Edwards, 2006; Witt and Edwards, 2000

Notes on Natural Enemies

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The following insect natural enemies were found feeding on A. asparagoides foliage during surveys in South Africa: larvae of moths Agrotis sp., Euplexia augens, Lycophotia oliveata and Cuculliaterrensis, an unidentified cecidomyiid gall fly, a Hespera sp. flea beetle, an undescribed Erythroneurini leafhopper (formerly referred to as Zygina sp.) (M Fletcher, NSW Agriculture, Australia, personal communication, 2009), a Crioceris sp. leaf beetle and adults of the weevil Brachycerus parilis (Kleinjan and Edwards, 2006). Larvae of the moth Zalaca snelleni  and the wasp Eurytoma sp. were also found feeding on fruits or immature seeds. No insects were observed feeding on below-ground organs despite the extensive excavations undertaken. The rust fungus Puccinia myrsiphylli was the sole pathogen observed attacking foliage of A. asparagoides (Kleinjan et al., 2004b).
 
The southwestern Cape form of A. asparagoides was also examined for natural enemies during surveys. Kleinjan and Edwards (2006) observed the Erythroneurini leafhopper, Zalaca snelleni and Eurytoma sp. on above-ground parts as well as weevil larvae, probably of a Brachycerus species, feeding on tubers. Infection by the rust fungus was never seen in the field on this form of A. asparagoides.
 
In the introduced range in Australia, very little leaf damage and no fruit or seed damage were observed during surveys of 34 sites infested by A. asparagoides in Western Australia in 1996–97, prior to the release of biological control agents (KL Batchelor and JK Scott, CSIRO Entomology, Australia, personal communication, 2010). In Victoria, the weevil Phlyctinus callosus and larvae of an unidentified Lepidoptera were observed feeding on foliage in the mid-1990s, while light brown apple moth (Epiphyas postvittana) larvae were seen eating ripe fruit pulp but not damaging seeds (Raymond, 1999). Damage by snails (Helix aspersa) has been observed from time to time in New South Wales.
 
In the introduced range in New Zealand, a survey of 32 sites between November 2005 and January 2007 identified 76 generalist herbivorous invertebrate species on A. asparagoides (Harman et al., 2008). The anthracnose fungus Colletotrichum gloeosporioides was isolated from necrosis on stems and cladodes, although damage overall was low. Several weak and opportunistic pathogens or saprophytic fungi were also isolated from minor damages.

 

Means of Movement and Dispersal

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Natural Dispersal (Non-Biotic)
 
Established plants can increase in size through spread of branching rhizomes bearing meristematic buds. Stansbury and Scott (1999) estimated from a landholder survey that patches of about 10 m2 expanded radially by approximately 0.6 m year-1. Situations where seeds were dispersed downstream by flood events have been observed (Graham and Mitchell, 1996).
 
Vector Transmission (Biotic)
 
Frugivorous birds are recognised as important contributors to seed dispersal in A. asparagoides (Stansbury, 1996, 1999, 2001; Raymond, 1999). At least 12 bird species, both native and exotic, have been recorded feeding on berries and potentially dispersing seeds in southern Australia (Stansbury, 2001). Ground-foraging birds, such as common blackbirds (Turdus merula) and emus (Dromaius novaehollandiae) also contribute to secondary dispersal of fruits (Loyn and French, 1991; Raymond, 1999). Bird dispersal of seeds is highly stochastic and consequently dispersal is difficult to predict precisely (Siderov et al., 2006). The dispersal pattern is influenced by the behaviour of different bird species and habitat structure, since many birds fly to perch-trees to consume fruits. Stansbury’s (2001) diffusion model showed that A. asparagoides distribution at Bold Park in Western Australia was correlated with the flight pattern of silvereyes, with a mean distance for seed dispersal of 90.5 m. Occasional long-distance dispersal is also possible and may significantly increase rate of spread at a landscape scale. Based on silvereyes’ gut passage rates and flight speed, the maximum potential seed dispersal distance was estimated to be 12 km. Gradual short-distance bird dispersal of seeds along roadsides and wildlife corridors that connect disparate habitats has the potential to increase the rate of invasion of remnant vegetation patches at a local scale (Siderov et al.,2006).
 
A. asparagoides seeds are also dispersed by other animal vectors, such as rabbit (Oryctolagus cuniculus) and foxes (Vulpes vulpes) (Graham and Mitchell, 1996; Raymond, 1999).
 
Accidental Introduction
 
Careless disposal of garden waste and earthworks (e.g., roadside grading) can spread rhizomes over considerable distances to start new infestations. Mud containing seeds that adheres to animals, vehicles and machinery is also believed to contribute to dispersal (Parsons and Cuthbertson, 2001), although this would be of relatively minor importance.
 
Intentional Introduction
 
Deliberate movements of plants or propagules within and between countries occurred widely in the past for ornamental purposes. Rhizome fragmentation has been used locally to increase numbers of plants for sale, although such sale is now prohibited in Australia.

 

Impact Summary

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

Economic Impact

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It is estimated that at least 20% of growers who manage a total of more than 6500 ha of citrus orchards in districts bordering the Murray River from Mildura to Cobram in Australia, are affected by A. asparagoides (Kwong and Holland-Clift, 2004). Grower’s perceptions of the negative impacts of A. asparagoides on their orchard ranged from increased labour and financial costs, damage to citrus trees, reduced fruit production, impediment to irrigation and fruit harvesting, and unsightliness. The cost of control is estimated to be as high as $2000 ha-1 y-1. In very severe infestations, where >80% of the orchard is infested, growers believed that it is often cheaper in the long run to remove trees and replant.

Environmental Impact

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Impact on Habitats
 
Rapid growth of A. asparagoides in autumn and its climbing habit provide canopy dominance and make it highly competitive. Once an infestation is established, the amount of light reaching the soil surface is very low, thereby preventing other plants from persisting. Raymond (1999) reported that the amount of irradiance reaching the soil surface was reduced by 93% under a canopy of A. asparagoides foliage. Competitiveness of A. asparagoides is also due to the quantity of below-ground biomass that occupies most of the space available in the substrate. During the fruiting stage, established infestations can harbour large numbers of exotic birds (Raymond,1999; Stansbury, 2001), which are considered pests in Australia.
 
A. asparagoides has been identified as a threat to four endangered ecological communities in New South Wales (Coutts-Smith and Downey, 2006). Eleven additional ecological communities, including six already listed under the New South Wales Threatened Species Conservation Act 1995, are potentially threatened by A. asparagoides (Downey, 2006).
 
Impact on Biodiversity
 
A. asparagoides is a highly competitive species that poses a direct threat to at least 17 native plant species in South Australia and New South Wales (Quarmby, 2006; Downey, 2006). In addition to the species listed in Downey (2006), the critically endangered orchid species Pterostylis bryophila and Pterostylis sp. ‘Halbury’ in South Australia are also considered under threat due to habitat loss and primarily A. asparagoides invasion (Quarmby, 2006). Furthermore, a recent assessment made in southeastern New South Wales, which involved an extensive consultation process, suggested that up to 52 additional plant species are potentially threatened by A. asparagoides invasion (Downey, 2006). The black grass-dart butterfly (Ocybadistes knightorum) is also listed as threatened by A. asparagoides in New South Wales (Coutts-Smith and Downey, 2006).

 

Threatened Species

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Threatened SpeciesConservation StatusWhere ThreatenedMechanismReferencesNotes
Beyeria subtectaNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Eremophila barbataNational list(s) National list(s)South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Leionema equestreNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Olearia microdiscaNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pimelea spicataNational list(s) National list(s)Australia; New South WalesCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pomaderris halmaturina subsp. halmaturinaNational list(s) National list(s)South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Prasophyllum pallidumNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Prostanthera calycinaNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pseudanthus micranthusNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pterostylis arenicolaNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pterostylis bryophilaCR (IUCN red list: Critically endangered) CR (IUCN red list: Critically endangered)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringQuarmby, 2006
Ptilotus beckerianusNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Pultenaea pedunculataNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Spyridium coactilifoliumNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Spyridium eriocephalumNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006
Thelymitra epipactoidesNational list(s) National list(s)Australia; South AustraliaCompetition - monopolizing resources; Competition - shading; Competition - smotheringDowney, 2006

Risk and Impact Factors

Top of page Invasiveness
  • Proved invasive outside its native range
  • Has a broad native range
  • Highly adaptable to different environments
  • Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
  • Pioneering in disturbed areas
  • Tolerant of shade
  • Long lived
  • Fast growing
  • Has high reproductive potential
  • Has propagules that can remain viable for more than one year
  • Reproduces asexually
  • Has high genetic variability
Impact outcomes
  • Altered trophic level
  • Damaged ecosystem services
  • Ecosystem change/ habitat alteration
  • Increases vulnerability to invasions
  • Modification of nutrient regime
  • Modification of successional patterns
  • Monoculture formation
  • Negatively impacts agriculture
  • Negatively impacts forestry
  • Reduced amenity values
  • Reduced native biodiversity
  • Threat to/ loss of endangered species
  • Threat to/ loss of native species
Impact mechanisms
  • Competition - monopolizing resources
  • Competition - shading
  • Competition - smothering
  • Rapid growth
Likelihood of entry/control
  • Highly likely to be transported internationally deliberately
  • Difficult to identify/detect in the field
  • Difficult/costly to control

Uses

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Economic Value
 
A. asparagoides was widely grown as an ornamental plant from the mid-1880s to the early twentieth century, but its popularity declined substantially afterwards. It is not sold anymore in nurseries in Australia and it is legislated against in all states and territories (Morin et al., 2006a).
 
Social Benefit
 
In South Africa, A. asparagoides is believed to have therapeutic attributes. A lotion prepared from its root biomass is used to bathe sore eyes (Guillarmod, 1971), although no detail on efficacy and extent of use is provided in the literature. Tubers have been seen being sold in the Cape Town market as a remedy for stomach problems, but again no detail is known of the recommended method of preparation or the dose rate (PB Edwards, CSIRO Biological Control Unit, University of Cape Town, South Africa, personal communication, 2006.
 
Environmental Services
 
A. asparagoides’ fruits form part of the diet of a small number of Australian native birds (Stansbury, 2001), whose normal host plants may occur in low density due to habitat degradation or destruction. Its foliage is non-toxic to mammals. It is reported to form a significant part of the diet of tammar wallabies (Macropus eugenii) on Garden Island in Western Australia (Bell et al., 1987). Sheep and cattle have also been observed grazing on the foliage (Siderov and Ainsworth, 2004), but there is no information on its nutritional value.

 

Uses List

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Environmental

  • Amenity
  • Landscape improvement

General

  • Botanical garden/zoo

Medicinal, pharmaceutical

  • Traditional/folklore

Ornamental

  • Cut flower
  • Potted plant
  • Propagation material

Detection and Inspection

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A. asparagoides is easily detected in the field during the growing season when extensive foliage is produced, but not necessarily at other time of the year after it has senesced and only dead stems remain. Prior to initiation of a control program, it is useful to map the distribution and densities of infestations in the targeted area or region (Carr, 1996). Such a map provides the basis for prioritising areas for control. The Asparagus Weeds Best Practice Management Manual (2006) recommends to thoroughly check tree corridors, roadside vegetation and tall trees on the verge of native vegetation areas for presence of plants. Searches should be performed up to several hundred metres beyond the location where the last plants are found to ensure that all bird-dispersed seedlings are located.

Similarities to Other Species/Conditions

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Following an extensive examination of herbarium specimens and field surveys in South Africa, Kleinjan and Edwards (1999) identified two distinct forms of A. asparagoides: a widespread form that corresponds to the invasive populations found across temperate Australia and another form that is restricted to the south-western part of the Western Cape Province of South Africa. The two forms may warrant separation at the species level. The south-western Cape form is differentiated from the widespread form by its rhizomes that grow primarily upwards towards the soil surface, larger tubers and more waxy and elongated cladodes. It was not reported from Australia before 2004, when a few small populations were located in southeastern South Australia and western Victoria (Coles et al., 2006). This form of A. asparagoides is less susceptible to the biological control agents released in Australia.

A. asparagoides is somewhat similar to other asparagus ferns (Asparagus spp.), however it can be easily distinguished from them by its larger and broader 'leaves' (10-70 mm long and 10-30 mm wide) (Queensland Government, 2012). In Australia, it is more often confused with many native vines that have similar leaves: distinguishing features are given in the A. asparagoides datasheet from the Queensland Government (2012).

Prevention and Control

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Prevention      
 
Public awareness
 
Since A. asparagoides was listed as a Weed of National Significance in Australia (Thorp and Lynch, 1999), major education campaigns have been undertaken to increase community awareness of the problems it causes and available methods of control (Weeds Australia, 2010).
 
Eradication
 
Eradication of A. asparagoides at a local scale is only feasible for recently established infestations and prior to plants producing fruits. A. asparagoides has been the target of an eradication campaign for several years in Tasmania (Cooper and Warren, 2006), but the campaign’s effectiveness is now being questioned as new small infestations keep on emerging. An eradication campaign was also initiated in 2004 on Lord Howe Island off the coast of New South Wales and is intended to last for a minimum of 15 years (Le Cussan, 2006).
 
Containment/zoning
 
Since most A. asparagoides seeds are bird-dispersed within 300–500 m of the source (Stansbury, 2001; Siderov et al., 2006), it is recommended to control outlying plants or patches within a buffer zone of 500 m around the edge of a main infestation (Asparagus Weeds Best Practice Management Manual, 2006). Siderov et al. (2006) emphasised the importance of controlling A. asparagoides along paths and tracks within remnant vegetation because they act as a conduit for invasion. Alternatively, reducing the number, or modifying the spatial arrangement, of tracks within a reserve can be considered to limit seed dispersal within the area.
 
Control
 
Carr (1996) recommended initially focusing control efforts on low density infestations occurring in bushland with the highest conservation value before tackling more severely infested areas. Targeting control towards climbing plants that produce the most fruits within the core infestation may also help limit spread.
 
Cultural control
 
Gardeners are encouraged to prevent further spread by avoiding composting, mulching or dumping garden refuse containing A. asparagoides rhizomes from their properties. Dug-up plants should be placed in black plastic bags and left out in the sun for many months to kill rhizomes (Asparagus Weeds Best Practice Management Manual, 2006). Hygiene of earth-moving equipment is important to prevent spread of viable rhizome fragments that may be present in soil (Graham and Mitchell, 1996).
 
Physical/mechanical control

Isolated young A. asparagoides plants with an underdeveloped root system can be pulled-out by hand when growing in loose soil (Carr, 1996). Manual removal of mature plants and their root system is only appropriate for small isolated infestations, particularly in areas of high conservation value (Carr, 1996; Asparagus Weeds Best Practice Management Manual, 2006). It is also used by growers in citrus orchards who are reluctant to use herbicides (Kwong and Holland-Clift, 2004). Follow-up control is often required because A. asparagoides regenerates from small pieces of living rhizome left behind in the soil. Removed root mats should be disposed of by deep burial (>2 m deep) or dried and burnt (Graham and Mitchell, 1996; Pike, 1996). Hand removing is time and labour intensive and consequently not suitable for severe and extensive infestations. Removal of well-developed mats of rhizomes and tubers can create excessive disturbance that hinders natural bush regeneration and facilitates colonisation by invasive plants.
 
Mechanical removal of above-ground biomass is effective at reducing seed production if performed before fruiting, but many repeated slashings are necessary to exhaust below-ground reserves. Slashing is sometimes carried out in citrus orchards around infested trees (Kwong and Holland-Clift, 2004). A. asparagoides is palatable to mammals (Bell et al., 1987; Carr, 1996; Siderov and Ainsworth, 2004) and thus livestock grazing is potentially an effective control method. Siderov and Ainsworth (2004) found no A. asparagoides occurring next to fenced areas of pasture where it was abundant and supported the implementation of controlled grazing in areas fenced off for revegetation to keep the invasion in check. Such an approach may be more efficient than regularly searching the area to remove newly-established seedlings, but may not be appropriate when the area is revegetated with native species palatable to livestock. Cooke and Robertson (1990) reported use of sheep to reduce A. asparagoides density prior to chemical control on Kangaroo Island, South Australia.
 
A. asparagoides is not killed by bushfires unless the fire is very intense. Hot summer fires can destroy a large proportion of rhizomes and tubers lying near the soil surface, but generally fire does not affect below-ground reserves located deeper in the soil (Carr, 1996; France, 1996). Land managers are encouraged to take advantage of a late summer or early autumn wild fire and implement chemical control since A. asparagoides is usually one of the first species to emerge in the burnt areas before regeneration of sensitive native species (Carr, 1996; France, 1996; Graham and Mitchell, 1996).
 
Biological control
 
Three biological control agents for A. asparagoides, all of South African origin, have been released in Australia: An undescribed Erythroneurini leaf hopper in 1999, the rust fungus Puccinia myrsiphylli in 2000 and a Crioceris sp. leaf beetle in 2002 (Witt and Edwards, 2000, 2002; Kleinjan et al., 2004b; Morin and Edwards, 2006; Morin et al., 2006b). A Eurytoma sp. seed wasp was also considered a promising candidate agent, but host-specificity testing could not be performed dueto rearing difficulties and concerns raised about the conflict of interest with producers of cultivated asparagus seed. Large-scale releases of the leaf hopper and rust fungus were carried out in partnership with community groups, land managers and schools (Kwong, 2002; Batchelor et al., 2004; Overton and Overton, 2006).
 
Impact of the agents on A. asparagoides has been assessed in a series of glasshouse and field experiments (Batchelor and Woodburn, 2002; Morinet al., 2002, 2006b; Kleinjan et al., 2004a; Spafford Jacob et al., 2007; Turneret al., 2008a, 2010). The rust fungus has had a major impact in reducing A. asparagoides populations, particularly in moist coastal areas of Australia where climatic conditions are conducive to epidemics. The leaf hopper has also adversely affected A. asparagoides at some sites, but its populations have a tendency to fluctuate from year to year, hence limiting impact. Establishment of the leaf beetle has been extremely poor despite considerable release efforts and therefore this agent is not currently contributing to control.
 
Puccinia myrsiphylli was found in New Zealand in November 2005 near Auckland, New Zealand, probably as a result of an accidental introduction or natural dissemination of spores from Australia on wind currents (Waipara et al., 2006). Subsequent surveys and field assessments revealed that it was widespread throughout the range of A. asparagoides in northern New Zealand, often causing severe damage and premature defoliation (Harman et al., 2008).
 
Chemical control
 
Several herbicide trials have been conducted in the last 20 years (Cooke and Robertson, 1990; Dixon, 1996; Pritchard, 1996, 2002), identifying glyphosate, metsulfuron methyl and some related sulfonylureas as the most effective non-selective systemic herbicides against A. asparagoides. To achieve best results with chemical control, a spray program of at least three years is recommended. Repeat applications of herbicides are essential to kill plants that are missed in the first year, any new regrowth and seedlings, and to have a significant impact on below-ground biomass. A longer control program with herbicide applied every 2–3 years, rather than annually, may be a more efficient use of time and resources without compromising outcomes.
 
Best control outcomes have been achieved when metsulfuron methyl or glyphosate was applied onto actively growing A. asparagoides in the early to late-flowering stage (Dixon, 1996; Pritchard, 2002). Good control was also obtained with applications made at the flower bud to green berry stage (Pritchard, 2002). Spraying before flowering is advocated by some to allow better translocation to the rhizome (Cooke and Robertson, 1990) and to prevent off-target damage to indigenous species that emerge in spring (Carr, 1996).
 
Chemical control of A. asparagoides growing amongst citrus foliage is not generally practiced because of the high risk of damage to trees (Kwong and Holland-Clift, 2004).
 
IPM
 
A combination of biological and chemical control with other management tactics such as fire may be necessary in some areas to tackle A. asparagoides more effectively (Willis et al., 2003). Access to infested areas for spraying is considerably improved after a fire because most woody shrubs have usually been destroyed. Willis et al. (2003) suggested use of fire in autumn, after most annual shoots have emerged, and post-fire application of herbicide on regrowth to maximize depletion of below-ground reserves. Turner and Virtue (2009) assessed the impact of herbicide used following wildfire that burnt an area of mallee vegetation in South Australia infested by A. asparagoides. Ten years later there was almost 4 times less shoots of A. asparagoides in herbicide treated plots when compared to plots that were untreated at the time of the fire. There is no contra-indication to use biological control for A. asparagoides post-fire and agents are most likely to recolonize sites if they are present in unburnt nearby areas (Morin et al.,2006b).
 
Ecosystem Restoration
 
Natural ecosystems may take many years to recover after an area is cleared of A. asparagoides (Turner and Virtue, 2006). Presence of bare ground after A. asparagoides is controlled increases the likelihood of re-invasion or colonisation by other invasive plants. Land managers can either rely on natural recovery processes or actively revegetate controlled sites with indigenous species. Pike (1996) observed natural regeneration of native plants, such as Hardenbergia comptoniana and Acacia pulchella in an area where A. asparagoides had been hand removed in Western Australia. However, Turner et al. (2008b) found that the exotic seed bank germinated more readily than the native species in areas that have been invaded by A. asparagoides and that additional restoration efforts would be needed after successful control.

Gaps in Knowledge/Research Needs

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No research has yet been carried out to devise a practical and effective strategy to integrate the different control methods available for A. asparagoides. Detailed investigations into the mechanisms of invasion of A. asparagoides, including impact on soil nutrients and microbial communities, are needed to guide restoration approaches post-control.

 

References

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Links to Websites

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WebsiteURLComment
CSIRO - Management and control of bridal creeperhttp://www.csiro.au/science/BridalCreeper.html

Organizations

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Australia: Commonwealth Scientific and Industrial Research Organiation - CSIRO, CSIRO Entomology GPO Box 1700, Canberra ACT 2601, http://www.csiro.au/org/Entomology.html

Australia: Weeds Australia, An Australian Weeds Committee National Iniative - National Portal, http://www.weeds.org.au/WoNS/bridalcreeper/

Contributors

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30/04/10 Original text by:

Louise Morin, CSIRO Entomology, Clunies Ross Street, Acton, GPO box 1700, Canberra, Australia

Distribution Maps

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