Spotted Lanternfly

Biology  |  |  Impacts  |  Detection  |  Management


Origin and Spread

The spotted lanternfly, Lycorma delicatula, is an invasive species to the United States, first discovered in Pennsylvania in 2014. It was originally from China and southern Asian countries such as India. It is likely to become a serious agricultural pest without natural enemies to keep populations low. It was accidentally introduced into South Korea in 2006 and has spread dramatically to become a major agricultural pest, especially for grape production.

The risk of spread in the northern US was once believed to be low due to cold winters. More recently, however, many eggs and newly hatched nymphs have survived the winter. While not yet in New York, the spotted lanternfly is on the border with Pennsylvania and if it does move into NY and become established, it has the potential to become a significant agricultural pest causing untold physical and economic damage.


Spotted lanternfly, Lycorma delicatula, adult.  [Photos: Holly Raguza,]

This map shows documented sightings of the spotted lanternfly since it was first found through Oct. 17, 2017. (Courtesy Pennsylvania Department of Agriculture



Spotted lanternfly eggs hatch as nymphs in April and May during the early hours of the day. The black bodied nymphs go through 4 growth phases (instars) before becoming a winged adult. Instars 1-3 have white spotted black bodies, while the 4th instar develops black and red mottling under the white spots. Adult lanternflies have grayish forewings with black spots; the hind wings are red and black spotted on the lower portion and grey and black with a bold white stripe on the upper portion. Adult females are about an inch in length; males are about 4/5ths of an inch. Adults are strong hoppers but weak fliers. In the fall, adults often shift to feeding the invasive Tree of Heaven (Ailanthus altissima). In late September until early winter, adults lay oothecas, or egg sacs, which house up to 30-50 brown seedlike eggs with a shiny, light orange/brown waxy coating. The egg sacs have a smooth, shiny surface. While lanternflies often lay eggs on Tree of Heaven, they will use any smooth, vertical surface including other smooth-barked trees, stones, vehicles, outdoor furniture and other manmade surfaces.  Eggs in Pennsylvania overwintered successfully in 2014; once eggs hatch the waxy coating is removed, leaving parallel lines of hatched egg sacs behind.

Lanternfly; female caudal View.  [Photo: Lawrence Barringer, Pa Dept. of Agriculture,]


Lanternfly juvenile stage.  [Photos: Lawrence Barringer, Pa Dept. of Agriculture,]


Both adults and nymphs use their piercing and sucking mouthparts to eat the phloem tissue of a wide variety of plants in order to obtain nutrients.  The insects also excrete a sugary fluid similar to aphid honeydew, which encourages mold and disease growth.  In the native range of the spotted lanternfly, these impacts does not normally kill host plants; absence of  natural predators, however, can lead to overinfestaton and cause sickness and death in infested plants. Overfeeding by the lanternflies can extract a damaging percentage of the plant’s nutrients, and the dripping sap from lanternfly feeding wounds combined with sugary lanternfly excretia can lead to mold and disease damage.

Spotted lanternflies feed on over 65 species of plants, preferring plants that have high sugar content and toxic metabolites. These include many agricultural species such as fruit vines (grapes), fruit trees (apples, cherries, peaches, pears, plums) and maple trees. Ornamental plants and forestry species including dogwoods, lilacs and pines are also susceptible. The spotted lanternfly has the potential to become a serious agricultural pest and stressor on natural systems.

Honeydew secretions building up at tree base. This is a sign of heavy lanternfly infestation. [Photo: Lawrence Barringer, PA Dept. of Agriculture,]

Bark damage done by lanternflies.  {Photo: Lawrence Barringer, PA Dept. of Agriculture,]



In the spring, look for the white-spotted black-bodied nymphs and white-spotted, mottled red-and-black fourth instar nymphs feeding on any of the wide variety of host species, both woody and non-woody.  In the summer, adults are visible while feeding. Indicators of lanternfly damage are seeping sap wounds on non-woody and woody species, and patches of blackened soil around the plant base.  Other insects and molds are also attracted by the sugary secretions caused by the sap and honey dew, including ants, bees and wasps. In the fall, brownish egg sacs can be seen plastered to tree trunks (especially Tree of Heaven) or other smooth surfaces such as stones, vehicles, farm equipment or outdoor furniture.

Lanternfly egg mass. [Photo: Holly Raguza,]



Spotted lanternfly management will vary based on whether the insect is found inside or outside existing quarantine areas.  To date, several towns in Pennsylvania including District, Earl, Hereford, Pike, Rockland and Washington and the boroughs of Bally and Bechtelsville are under quarantine. Quarantine areas may expand if the spotted lanternfly is found elsewhere in the US.

Within quarantine areas:

Egg sacs should be scraped off the host surface, soaked in alcohol or hand sanitizer and thrown away.

Lesser Celandine

Origin & Distribution  |  Identification  |  Biology  |  Impacts  |  Control



The spring ephemeral lesser celandine (Ficaria    verna  , previously Ranunculus ficaria L.), also called fig buttercup, is an herbaceous perennial found throughout the northeastern United States. It prefers moist, sandy soil, and thrives along stream and river banks, in open forested flood plains, and in other wetland sites. It will grow in drier habitats, however, and as a garden escape can often be found in urban and semi-urban areas.

Lesser Celandine, a.k.a. Fig buttercup (Photo: David Nicholls, National Biodiversity Network Trust [NBN Trust], UK)

Origin and Distribution

Native to Europe, Asia and northern Africa, lesser celandine was likely introduced as an ornamental plant. The earliest herbarium specimen dates to 1867 from Pennsylvania. Lesser celandine has been reported throughout the northeastern United States and west to Missouri, and in the Pacific Northwest.

Lesser Celandine US distribution (Map: EDDMapS. 2015. Early Detection & Distribution Mapping System. The University of Georgia – Center for Invasive Species and Ecosystem Health. Available online at; accessed October 1, 2015.

New York State distribution of Lesser Celandine (Map: Accessed: Oct. 1, 2015)



Lesser celandine is low growing and mat-forming, with long stalked leaved densely arranged in a basal rosette. Mature rosettes can reach up to 30 cm (12 in) diameter and up to 30 cm (12 in) tall. The kidney shaped leaves are 4 to 9 cm (1.6 to 3.5 in) wide, smooth, and have wavy edges. Leaf stalks (petioles) are deeply grooved and U-shaped in cross section. Flowers are yellow, 2 to 6 cm (0.8 to 2.4 in) wide and have long (10 to 30 cm; 4 to 12 in) peduncles. There are five sub-species of lesser celandine, distinguished largely by reproductive characters. Ornamental varieties may have a range of flower colors.


Close-up of flower petals and stamens1;  Bulbils formed at end of mature petioles allow for self-propagation2:  Tubrous roots3

(Photos: 1 David Nicholls, National Biodiversity Network Trust [NBN Trust], UKAntonino Messina, Flora of North America; 3 Andrea Moro, Dipartimento di Scienze della Vita, Università di Trieste) 


Lesser celandine is ephemeral (short-lived), and its emergence is triggered by increased light availability in the early spring. Shoots emerge from late-March to mid-April depending on environmental conditions, and flowering, which may be linked to water availability, occurs from late April to mid-May. Some sub-species produce pale aerial bulbils in the leaf axils. Seed production occurs in late spring, and by summer the above ground vegetation dies back and the plant becomes dormant. Seeds do not mature on the plant; rather, they require an after-ripening period to fully mature.  Some varieties do not produce many viable seeds, and the aerial bulbils and tuberous roots are the primary mode of reproduction. Bulbils are dispersed through animal and environmental disturbance, and can be carried along waterways.

Lesser celandine plant structure (Illustration: Duna-Ipoly Nemzeti Park Igazgatosag, Hungary, at


Lesser celandine emerges earlier than most native plants, and may inhibit the development and reproduction of other spring ephemerals, particularly smaller species like spring beauties (Claytonia spp.), trilliums (Trillium spp.), and bloodroot (Sanguinaria canadensis). These native wildflowers are a source of nectar for bees and other insects in the early spring. The bare ground left behind after lesser celandine senesces in late spring may be colonized by other weedy species.

Lesser celandine taking over a forest understory. (Photo: Les Merhoff,

Control, and Management

Due to its ephemeral nature and capacity for vegetative reproduction, lesser celandine can be challenging to effectively control. Small infestations can be removed manually, though care must be taken to completely remove the tubers from the soil. Repeated, early season mowing may reduce or stay growth of lesser celandine, but risks dispersing aerial bulbils. Glyphosate based herbicides may also be effective, and will require far less soil disturbance than manual removal. A 1.5% concentration of a 39% to 41% glyphosate isopropyl-amine salt solution with a non-ionic surfactant is effective for spot applications. Herbicide is most effective when applied early in the season and, when possible, applications should be made before any non-target plants have emerged. Note: Always check state/provincial and local regulations for the most up-to-date information regarding permits for control methods. Follow all label instructions. Mention of chemicals in this profile does not represent a recommendation by NY Sea Grant or Cornell University.

Yellow Flag Iris

Origin  |  Biology  |  Description  |  Impacts  |  Control


Commonly grown and transplanted for its showy yellow flowers, yellow flag iris (yellow flag) has invaded wetlands and other aquatic and semi-aquatic habitats. Yellow flag can be found at the edges of streams and ponds, in open and forested flood plains, along shorelines, and in freshwater and brackish marshes.

Yellow flag iris, Iris pseudacorus. (Photo: J.S. Peterson. USDA NRCS National Plant Data Center (NPDC), on PLANTS Database)

Origin and Expansion

Yellow flag is native to temperate regions of Europe, Asia, and northern Africa. It was imported to North America as an ornamental plant as early as the late-1700s. The plant has since been deliberately propagated as a horticultural plant and for erosion control and in sewage treatment ponds. It is now present in all but four states.

North American distribution of Yellow Flag Iris (Source: EDDMapS. 2015. Early Detection & Distribution Mapping System. The University of Georgia – Center for Invasive Species and Ecosystem Health,; accessed September 29, 2015).

New York distribution of Yellow flag Iris. Yellow squares are approximated locations; red dots are accurate locations. (Map: accessed Sept. 2015)



Yellow flag is a perennial. Shoot emergence and most seedling germination occur in spring, though in mild winters shoots may survive and remain green throughout the year. Flowering begins by late-May and continues into early-July. Plants generally do not flower until the third year of growth. Flowers are pollinated by bees and a few species of long-tongued flies. Seed production occurs from August through October; each plant can produce several hundred seeds. Seeds are mainly dispersed by currents, containing an air pocket to help keep them afloat, and are capable of remaining afloat for more than a year. Seedlings germinate and establish best in moist but not waterlogged soil. Yellow flag expands through rhizome growth. The thick rhizomes can persist for over ten years in the soil and can survive for more than three months if dried. The rhizomes of old plants older than ten years often break into fragments, which may then be dispersed by water.



The sword-like leaves are flat, erect and linear with a raised midrib. The dark to blue-green blades are 25-90 cm long and have sharply pointed tips. The flowering stems are usually similar in size to the leaves (50-100 cm in length). Flowers are pale to bright yellow or cream colored and 7-9 cm wide. The large (4-8 cm) seed pod is 3-sided and angular and turns from glossy green to brown as it ripens. Each pod contains dozens of seeds densely arranged in 3 rows. Roots are 10-30 cm in length, and the fleshy rhizomes are 1-4 cm in diameter.


Close-up of Yellow Flag Iris flower (Photo: Nancy Loewenstein, Auburn University, Yellow Flag Iris seeds (Photo: Leslie J. Mehrhoff, University of Connecticut,


Yellow flag expands quickly via rhizomes, and can form dense monotypic stands that can replace and crowd out valuable aquatic plants like cattails and other, native, irises. The root system forms a dense mat which compacts soil and inhibits seed germination of other plants. Large yellow iris populations may also reduce the habitat available to native fish and waterfowl. Thick growths of yellow flag can clog irrigation systems and streams and, by trapping sediment in the roots, can narrow waterways. All parts of the plant are toxic to livestock and other animals.

Dense growth of Yellow Flag Iris (photo: Todd Pfeiffer, Klamath County Weed Control,


Control, and Management

Small clumps can be dug out, though this is only effective if the rhizomes are entirely removed. Mowed plants will regenerate from the rhizomes, so plants must be cut multiple times to exhaust their energy reserves. The sap may cause skin irritation, so gloves should be worn when handling cut or otherwise damaged stems. Glyphosate herbicides approved for aquatic use can be effective, particularly if applied to recently cut foliage and stems. No biological control agents have been released to control yellow flag.  [Note: Always check state/provincial and local regulations for the most up-to-date information regarding permits for control methods. Follow all label instructions.]

Japanese Virgin’s Bower

Identification  |  Look-alikes  |  Biology & Habitat  |  Prevention and Control



Japanese virgin’s bower (Clematis terniflora DC, also called sweet autumn clematis) is an invasive ornamental vine in the Clematis genus. It is widely planted in gardens for its profusion of small, showy white flowers in late summer and equally showy silvery, corkscrew, feathery seedheads. Given support it can reach 30’ in height.  Its prolific nature and aggressive growth make it a problem in natural areas, where it can smother mature trees and displace native vegetation. Japanese virgin’s bower is on New York State’s regulated species list; it can be transported, purchased, sold and planted, but it can only be planted where it will not threaten natural areas, and must be displayed for sale with signs explaining that it is an invasive species.



Japanese virgin’s bower is native to Japan and China.  It was brought to North America as an ornamental species.  Leaves are opposite and compound, with three to five heart-shaped leaflets 2-3½” long; the surfaces are dark green and shiny, paler on the undersides. It climbs using tendril-like petioles to cling to support. Flowers are about 1” across, white, four-petaled, and held in clusters of 3-15; the plant blooms in late summer to fall. Seeds are small and brown, with showy, silvery-grey, feathery, curving seedheads.



Japanese virgin’s bower looks like many other Clematis genus species; the closest match is a native plant also commonly called virgin’s bower, Clematis virginiana.  Flowers and seeds are similar between the two species, but leaves of C. virginiana are toothed and lobed as opposed to C. terniflora’s entire, ovate leaves. The other Clematis have larger flowers of various colors; these ornamental species and hybrids are not particularly aggressive.



Japanese virgin’s bower can survive a wide range of soil types and light availability, although it is most productive with shade on its roots and full sun on its foliage. It establishes in hedgerows, waste spaces, roadsides, stream banks and forest edges. It is a semi-woody perennial, producing foliage on new stems each year. In southern climates it is semi-evergreen. Japanese virgin’s bower produces shoots from last year’s woody stems in spring. It flowers in August and produces seeds by late September, which are dispersed by wind.



Cultural: The similar native virgin’s bower, Clematis virginiana, has many of the same ornamental characteristics as Japanese virgin’s bower and is less aggressive.  They are so similar they are sometimes sold interchangeably at nurseries. Planting a native or noninvasive alternative will reduce the chances of this invasive damaging nearby natural areas.

Monitoring: Japanese virgin’s bower is most easily seen in the late summer, when it produces a froth of white flowers. Otherwise, look for a vine with small, oval- to heart-shaped, opposite leaves.  There is currently no research on control of this species; all information is from gardening and extension sources.

Manual and Mechanical Control: Remove the plants and its roots. Some root fragments may resprout, so repeat visits may be necessary. Given the plant’s prolific nature, expect seeds still in the soil to continue to sprout new plants for some time. Research on the longevity of C. terniflora’s seeds in the soil is not available.

Biological Controls: Currently there are no biological controls for Japanese virgin’s bower.

Chemical Controls: Japanese virgin’s bower responds to control with glyphosate and triclopyr and other common broadleaf herbicides, with no known resistance issues. However, all will require repeat treatments (Langeland and Meisenburg, 2009). Before using chemical controls, always check with your state’s environmental regulatory agency to check on the legality of using any particular chemical.


Walnut Twig Beetle, Thousand Cankers Disease

Distribution  |  ID & Biology  |  Hosts & Habitats  |  Impacts  |  Prevention & Control


The walnut twig beetle, Pityophthorus juglandis, is a tiny Scolytid bark beetle native to the southwestern US. It is the insect vector of Thousand Cankers Disease. The disease, first observed in the 1990s but not recognized until 2008, has killed many walnut trees planted outside their native range across the western US.  It is now a serious threat to walnuts in their native eastern range.


The walnut twig beetle was first detected in the Eastern US in 2010 and has now been found in Tennessee, Virginia, Pennsylvania, North Carolina, Ohio, and Maryland. It is widespread across Western US. While not yet in New York, the walnut twig beetle has been confirmed in adjacent states and poses a threat to walnuts in New York, the Northeast and New England.


Map Source:


Identification and Biology

The walnut twig beetle is reddish brown to brown and only 1.5-2mm long. Identification of this tiny bark beetle is possible with a dissecting microscope but confirmation by an expert is recommended because there are many similar species.  A very good identification guide is “LaBonte and Rabaglia, A Screening Aid for the Identification of the Walnut Twig Beetle, Pityophthorus juglandis Blackman.” The fungus is carried into the tree on the bodies of adult beetles; the beetles bore under the bark, create tunnels or galleries, and lay their eggs.


Steven Valley, Oregon Department of Agriculture,

Steven Valley, Oregon Department of Agriculture,

Hosts and Habitats

All walnut species tested so far are susceptible, particularly black walnut but also the Northern and Southern California black walnut species.


Thousand Cankers Disease, caused by a fungus and spread by the walnut twig beetle, is usually lethal to walnut trees (Juglans spp), particularly black walnuts (Juglans nigra), within a few years of noticeable symptoms. Black walnut has high economic value for wood production. Throughout its native range, the net volume of black walnut growing stock on timber land was valued at over $500 billion.


          Evidence of Thousand Cankers Disease. Ned Tisserat, Colorado State University,

Tree Killed by Thousand Cankers Disease.  Karen Snover-Clift, Cornell University,

Prevention and Control

Prevention through quarantine is key because symptoms (thinning canopy, leaf yellowing/wilt, tip die-back) appear after it is too late to save infected trees or nearby trees.  Most eastern states where the disease has been detected have adopted quarantines on the movement of nursery walnuts and walnut wood. Maps of range and quarantine information can be found here.

Heat treatment of logs before movement for processing has been shown to be an effective way to prevent spread of both the disease and beetle.

Pheromone baited traps have been developed for beetle detection in new areas but are only effective at short range.

Giant African Land Snail

Range & Distribution  |  ID & Biology Hosts & Habitat  |  Impacts  |  Prevention & Control


The giant African land snail was originally introduced to Hawaii in 1936 and Florida in 1966. Florida’s original eradication campaign took ten years and cost one million dollars. The snail was rediscovered in 2011. Eradication efforts are ongoing (2015).

Giant African land snails are eaten in many countries and sold as canned pet food for skinks, turtles, monitors, and small animals.


Giant African land snails, Lissachatina fulica, can grow up to 8 inches (20 cm) long. (Photo: Andrew Derksen, FDACS/DPI,


Yuri Yashin,,
Yuri Yashin,,

Range and Distribution

Giant African land snails are native to East Africa and found in Asia. In the USA, in Southern Florida and Hawaii, the snails are under quarantine. The USDA APHIS (Animal and Plant Health Inspection Service) has established additional regulated areas in Florida (June 2015). The snails are sold and raised as pets in other countries, including those of Europe. While not yet in New York, the giant African land snail, owing to the illegal pet trade, is prohibited in the state.


Identification and Biology

One of the largest terrestrial snails, full-grown adults can reach almost 8 inches (20 cm) long and 5 inches (13 cm) in diameter.  Adult shells are brownish with darker brown lengthwise stripes, have seven to nine whorls including a swollen long body whorl, and covers at least half the length of the snail. Snails have female and male reproductive organs. One mating can result in multiple clutches of eggs over time. Rapid population increases are likely because each snail can produce 1,200 eggs per year.

Close-up of giant African land snail. (Photo: Yuri Yashin,,

Giant African land snail egg clutch. (Photo: Yuri Yashin,,

Hosts and Habitats

The snails are found in many plant habitats and are known to preferentially consume beans, peas, cucumbers, melons, and peanuts.  Also at risk are ornamental plants, tree bark, and even the plaster, stucco, or paint on buildings.

No surface is off-limits to the snails. Giant African land snail on a Florida refuse bin. (Photo: Andrew Derksen, FDACS/DPI,


Because of their large size, ability to consume over 500 different kinds of plants, and cause damage to plaster and stucco buildings, the giant African land snail is one of the most damaging snails in the world. The snails are also a potential risk to human health because they can carry a parasitic nematode that can cause meningitis.

Giant African land snail infestation in Florida tree. (Photo: David G. Robinson, USDA APHIS PPQ,

Prevention and Control

Giant African land snails are able to survive cold temperatures in a semi-hibernation state. They represent a potential threat to New York even though they thrive in tropical/subtropical areas. If a snail shell is larger than two inches (5-6 cm) it is most likely a type of giant snail. Do not handle with bare hands. Importation is prohibited and specimens will be confiscated by customs. Do not purchase as pets or as educational animals through foreign online dealers or local distributors. For safe removal, or if found outdoors or for sale, contact local New York Department of Environmental Conservation, Cornell Cooperative Extension, or USDA offices.



Introduction  |  Biology and Habitat  |  Identification  |  Similar Species  |  Ecological Impacts  |  Control



Wineberry is an invasive shrub in the same genus (Rubus) as raspberries and blackberries. Wineberry creates spiny, inpenetrable thickets that reduce an area’s value for wildlife habitat and recreation. It was introduced to North America in the 1890s as breeding stock for raspberries. It was found invading natural areas by the 1970s, and it is currently recorded in most states east of the Mississippi River and in Alabama (USDA PLANTS Database). Wineberry replaces native vegetation, including native edible berry shrubs. It is differentiated from other berry-producing canes by the reddish appearance of its stems (caused by a dense coat of red hairs), silvery underleaf surfaces, and bright red berries.  Management can be obtained through mechanical, chemical, or combination of control methods.


Wineberry flowers. (Photo: Leslie J. Mehrhoff, University of Connecticut,

U.S. range of wineberry. (Map: USDA NRCS PlantsDatabase)

New York range of wineberry (Map: Accessed: Oct. 21, 2015)

Biology and Habitat

Wineberry is a close relative of other raspberries and blackberries. It grows in long shoots called canes up to six feet long, which can re-root at the tips when they touch the ground. Wineberry canes grow in two stages; in the first year they form a vegetative cane, and in the second year the cane becomes woody and produces lateral branches, flowers, and fruit (technically drupes, an aggregation of single seeded drupelets, but for clarity the term fruit will be used). Wineberries are perennial; while the canes each live two years, the plant produces new canes every year. Leaves are produced in April, flowers in May, and fruit from late June to August; leaves drop in late November. Wineberry does not need pollen from another individual to set seed, and therefore may reproduce more easily than natives like saw-toothed blackberry (Foss 2005).

Wineberry has a wide range of tolerance for light, soil type, and moisture level, and is hardy to USDA Zone 5 (annual minimum temperatures to -20F). While it is most productive in edge and wasteland habitats, it can be found in most habitats that exist in New York (Innes 2009), including forested habitats. Wineberry seeds are spread by animals, and seeds dropped on the forest floor can germinate when falling trees provide light to the forest floor. Once established, wineberry can persist indefinitely and reproduce once further disturbance occurs (Innis, 2005).

Wineberry canes. (Photo: Leslie J. Mehrhoff, University of Connecticut,



Wineberry is related to other raspberries and blackberries, and shares characteristics of both. Like raspberries, wineberry has silvery underleaves, a fruit core that remains on the stem when the ripe fruit is picked, and thorns. It is differentiated from other raspberry species by the fine red hairs that grow densely on its stems (and flowers) causing a reddish hue to the plant. Wineberry fruit is vibrantly red when ripe, which helps differentiate it from native black raspberries and blackberries; it also has three leaflets per leaf rather than five, which separates it from many blackberry species. Unique to wineberry is its small, greenish, hairy flowers with white petals and the way its fruit remain covered by sepals (greenish petal-like structures) until almost ripe.

Wineberry stem hairs. (Photo: Leslie J. Mehrhoff, University of Connecticut,

Top and bottom of wineberry leaves. (Photo: Leslie J. Mehrhoff, University of Connecticut.

Wineberry hairs. (Photo: Ansel Oomen,

There is quite a range of native and introduced Rubus species in New York; the wikibook Flora of New York has an excellent, easy to navigate identification website which includes wineberry. The more common species easily confused with wineberry are shown below. For a more complete look at the Rubus genus, the Flora of Michigan has an excellent online key.


Similar Species:

Rubus odoratus (purple-flowering raspberry or thimbleberry) has maple-shaped leaves that are soft and hairy; leaves not silvery; flowers pinkish-purple. Fruit is flatter and fuzzier than a raspberry, forming more of a cup shape.

Black raspberry (Rubus occidentalis) has whitish underleaves, but flowers hold their white petals out from the center of the flower, and fruit are usually purple-black (occasionally golden). Stems are green with a bluish cast that rubs off and have sparse, fairly robust thorns. Canes tip-root.

Red raspberries (Rubus idaeus, Rubus strigosus and many hybrids) have whitish underleaves and white petals, with red fruit, like wineberries. Stems are not covered in red hairs, are more lightly armed than black raspberry, and lack the bluish-white cast on their stems. Flowers might have a few hairs, but are not densely hairy like those of wineberry.


Rubus idaeus (red raspberry) cane. (Photo: Glen Mittelhauser.    Red raspberry fruit. (Photo: Alan Cressler)

New York has several species of native blackberries, all of which have green rather than silvery underleaves and solid-cored fruit (mostly black when ripe). Some have five to seven leaflets. Identification to species can be difficult. While the skin on some species is reddish or purplish, none are covered in reddish hairs like wineberry, and many are heavily armored with thorns.

Black raspberry (Rubus occidentalis) leaves and canes. (Photo: D. Cameron, from Go Botany website:

Evergreen blackberry (Rubus laciniatus) is an invasive blackberry. It has highly dissected leaves and black fruit with a solid core.

Evergreen blackberry canes and leaves. (Photo: Joseph M. DiTomaso, UC Davis.

Evergreen blackberry leaves and unripened fruit. (Photo by Joseph M. DiTomaso, University of California – Davis,

Himalayan blackberry (Rubus armeniacus) is also an invasive blackberry. It has stout, heavily armed but not hairy stems that grow up to 20 feet, tip roots like wineberry does, and produced large, sweet, dark-purple to black solid-cored fruit. It is the only blackberry with a whitish or grey-green underleaf, but usually has five leaflets instead of three, which along with its pinkish-white flowers and black fruit differentiate it from wineberry.

Himalayan Blackberry canes. (Photo: Joeph M. DiTomaso, UC Davis.

Ecological  Impacts

Wineberry can form dense, impenetrable thickets in natural areas, making the habitat unusable for some species and creating hiding places for others. It is more aggressive than many of the native raspberry and blackberry species, and has a wider range of tolerance for light, soil type, and moisture. Its establishment in forest understories as disturbance occurs can lead to its spread even in mature forests. There has been no study to date documenting its specific impact on native species.

Wineberry plants choking understory of second growth forest. (Photo: John M. Randall, The Nature Conservancy.


Wineberry control is more straightforward than control of many other invasive plants in New York. While any root fragments may start a new plant, wineberry does not have a vigorous underground storage structure; this makes it easier to control than, for instance, Japanese knotweed or lesser celandine. It is also susceptible to common pesticides.

For any invasive species control project, it is important to have a plan for the location before control begins.  Disturbance without replanting often results in the return of either the same invasive species or other invasives to the site; have a restoration plan in place before starting invasive species removal.

Mechanical control

Hand pulling wineberry or digging with a spading fork can be a successful strategy in small patches or where repeat visits are not costly, particularly if native species are planted where the ground has been disturbed. Return visits for a few years will be necessary to remove new plants that sprout from root fragments. As wineberry is armed with thorns and hairs, minimizing exposed skin during mechanical control is advisable.

Chemical control

Wineberry can be controlled using systemic herbicides such as glyphosate or triclopyr (Bargeron et. al., 2003). When using pesticides, be aware that many pesticides are prohibited within 100’ of water, as they are toxic to aquatic life and/or fail to break down in water. Some formulations of glyphosate-based herbicides are permitted for use near water, but the most common formulation (Roundup) is not permitted for use near water due to an adjuvant (chemical that helps the glyphosate stick to plant surfaces) that is toxic in aquatic habitats. Triclopyr also has both aquatic-permitted and prohibited formulations; choose carefully based on the characteristics of your treatment area. Always follow instructions on the label of any pesticide, and remember that New York has its own regulations for pesticides, both for the entire state and for specific regions like Long Island that have special environmental considerations. For New York State regulations, visit the DEC website:

Foliar application and cut-stump application are both recommended in various fact sheets (Massachusetts Audubon, Innes 2009, bugwoodwiki), but no experiments have been published on the relative efficacy of pesticides or application methods on wineberry (2015).


Kudzu (Pueraria montana)

Introduction & Distribution  |  Biology & Identification  |  Habitat & Ecology  |  Impacts  |  Control  |  Policy


Kudzu, “the vine that ate the South.” Kerry Britton, USDA Forest Service,


Kudzu is a semi-woody, trailing or climbing, perennial invasive vine native to China, Japan, and the Indian subcontinent. Kudzu is also known as foot-a-night vine, Japanese arrowroot, Ko-hemp, and “the vine that ate the South.” The vine, a legume, is a member of the bean family. It was first introduced to North America in 1876 in the Japanese pavilion at the Philadelphia Centennial Exposition. A second major promotion of kudzu came in 1884 in the Japanese pavilion at the New Orleans Exposition. The first recorded use of kudzu in North America was as a shade plant on porches in the American South (the plant produces attractive, fragrant purplish flowers in mid-summer). Kudzu was heavily promoted in the early-1900s when the government paid farmers to use the vine for erosion control (more than a million acres are estimated to have been planted as a result) and as a drought-tolerant, nitrogen-fixing legume (capable of bacterial growth with stem and root nodules converting free nitrogen to nitrates, which the host plant utilizes for its growth in low nitrogen soils) for livestock feed. During the Great Depression, thousands of acres of kudzu were planted by the Civilian Conservation Corps for hillside stabilization projects. In some areas, kudzu blossoms have been prized for their use in making kudzu blossom jelly and jam. The long kudzu fibers are also used in basket making. Ko-hemp, a more refined version of kudzu fiber has long been used for cloth weaving in China.


Use of kudzu for cattle grazing in the early-1900s. USDA NRCS Archive, USDA NRCS,


These government-sanctioned uses of the vine, combined with its innate, aggressive, range-expansion capabilities resulted in a rapid spread of kudzu throughout North America. Kudzu can now be found in 30 states from Oregon and Washington State to Massachusetts, particularly infesting states from Nebraska and Texas eastward most heavily; the vine is most common in the South. It has also been discovered in Hawaii and the warm, south-facing growing region on the north shore of Lake Erie in the Canadian Province of Ontario.


U.S. range of kudzu. USDA PLANTS database, July 2014.



Kudzu is an herbaceous to semi-woody, climbing or trailing, nonnative, deciduous, perennial vine or liana (a vine that is rooted in ground-level soil and uses trees and other vertical supports (telephone polls, buildings, etc.) to climb to the forest canopy to get access to light. A well-known example would be common wild grape).


Kudzu covering other vegetation and forming a liana. James H. Miller, USDA Forest Service,

Close-up of a kudzu liana. Leslie J. Mehrhoff, University of Connecticut,


Kudzu produces long, hairy vines from a central root crown. Kudzu has dark-green, hairy, alternate, compound leaves, 2 – 8 inches (5 – 20 cm) in length with three oval- to heart-shaped leaflets 3 – 4 inches (8 – 10 cm) long at the end; these leaves may be slightly or entirely lobed. Stems are also hairy. Vines can grow up to 30 to 100 feet (9 – 30.5 meters) per year. The vines have 0.8 – 1 inch (2 – 2.5 cm) flowers on 4 – 8-inch (10 – 20 cm) axillary racemes (short, equal length stalks along a main stem forming clusters of flowers with the oldest flowers toward the base with the newest end of the stalk terminating in one or more undeveloped buds). Vertical kudzu vines in full sunlight produce flowers in late-summer; horizontal vines seldom produce flowers. The flowers are typically red, purple, or magenta with a strong, grape-like aroma; pink or white flowers occur occasionally.


Kudzu leaflets found at end of stem. James H. Miller, USDA Forest Service,

Kudzu flowers made the plant popular for planting around porches. Forest and Kim Starr, Starr Environmental,


Population Expansion

Kudzu populations spread both asexually and by seed germination.

Asexual (vegetative) spread:

The most common method of spread is by setting new root crowns at almost every node where horizontal trailing stems come in contact with bare soil (this can be every few feet); new vines will form at these nodes the following spring and will spread out in all available directions. Kudzu tap roots can grow up to 12 feet (3.6 meters) long and weigh up to several hundred pounds. This may help kudzu to withstand long periods of drought.


A typical mature kudzu root crown. The Coalition To Control Kudzu Without Chemicals


Sexual spread:

Kudzu usually does not flower until its third year, with flowers and seeds forming only on vertical climbing vines. Kudzu produces clusters of 20 – 30 hairy brown seed pods, 1.6 – 2 inch (4 – 5 cm) long pods. Each pod contains from 3 to 10 kidney bean-shaped seeds, of which only 1 or 2 seeds are viable. Dormant  viable seeds are unable to germinate until after their seed coats have become water permeable as a result of physical scarification (breaking the seed coat by abrasion or prolonged thermal stress). Seeds deposited below the vines in the seed bank may take several years to germinate. This can be problematic during control efforts because it can result in the reemergence of the plants years after eradication was believed to have been achieved. It has been observed that kudzu in North America is more likely to grow asexually than by setting seed. It appears that this is due to kudzu seedlings being outcompeted by vegetatively produced vines.


A small cluster of kudzu pods. James H. Miller & Ted Bodner, Southern Weed Science Society,



Factors that help determine how invasive kudzu will be in any habitat appear to be climate and availability of light. Warmth and humidity are important factors, with greater colonization corresponding to warmer average annual temperatures and higher average humidity. To reach additional light, the vines climb existing vegetation and hard vertical surfaces. It does not appear that the composition of the local native plant community has much influence on kudzu invasiveness. Even undisturbed plant communities adjacent to an existing population of kudzu can be at risk. Typical kudzu habitats are usually open, disturbed areas such as roadside ditches, rights-of-way, and abandoned fields. In such settings, kudzu can form large monocultures with thousands of plants per acre.

Kudzu has a strong daily leaf orientation capability; by controlling the leaf position as it faces toward or away from the sun, kudzu can control sunlight intensity on the leaflets that are exposed. This ability can reduce leaf temperatures relative to native vegetation and minimize the amount of water lost from the plant by leaf surface transpiration during times of peak sunlight. It may also be a benefit below forest canopies where light is dim by increasing the surface area of leaves receiving sunlight. Leaves exposed to open sunlight may be able to maximize photosynthesis, store additional food in kudzu’s rhizomes, and have a competitive advantage over native vegetation.

Kudzu accumulates and maintains substantial carbon reserves in large woody, tuberous roots, again giving it a competitive advantage.

Trailing stems in open areas tend to die back in the winter. Vertically climbing vines develop thick bark and can reach diameters greater than 0.8 inch (2 cm), aiding in overwintering.

Kudzu vines can more easily grow around smaller vines such as honeysuckle (Lonicera spp.) than around bare tree trunks. This growth tactic appears to aid the plant in the formation of lianas in forested areas. Once established, kudzu lianas compete with forest trees both for sunlight in the crown and for water and nutrients from the soil. The vines may directly damage colonized trees by strangulation. These physical traits of a kudzu liana significantly impact the ability of native trees to grow and reproduce, increasing the early mortality of native trees, and preventing the establishment of new trees or shrubs in the dim light below the colonized canopy.

Kudzu lianas can cause weakened trees to fall from the weight of the overgrowth of vines or by pulling down trees attached to the liana when one weak tree succumbs to the weight of ice freezing onto the tree and/or the vines.

Kudzu thrives where the climate favors mild winters (40 – 60°F {4 -16°C}), summer temperatures rising above 80°F (27°C), precipitation greater than 40 inches (101 cm), and a long growing season.

Because of its underground root crowns, kudzu can escape fire damage. During the growing season, kudzu’s underground root system can provide significant water to the foliage; the high water content stems and foliage are able to resist some fire damage that may kill nearby native plants.

There is some indication (not yet definitively proven) that wildfire (or controlled burn) soil heating may promote kudzu seed germination by scarifying the seedcoat which would allow penetration by water to allow for germination.

Robert L. Anderson, USDA Forest Service,



Native Plant Community Impacts:

A kudzu invasion can cause several different types of major impacts on native plant communities: it can crowd them out; it can outcompete them; and it can physically crush them.

Since kudzu can fix nitrogen in its roots, it can thrive in soils too low in nitrogen to support robust growth of native vegetation, thereby outcompeting native plants for both nutrition and growing space, ultimately forming monospecific plant communities. This significantly alters natural plant communities and the animals that rely on those natural communities for food and habitat. Areas of more than 100 acres (40 hectares) with 1 – 2 plants per square foot, or 40,000 to 85,000 plants per acre (107,000 to 215,000 plants per hectare) can be found in the American South.

Kudzu’s rapid growth rate and its manner of growing over whatever it encounters in its path can also overwhelm native plant communities, also resulting in monospecific stands of the vine.

As heavy infestations of kudzu can completely cover trees of almost any size, kudzu lianas can both fell trees from their extreme weight or nearly eliminate light availability within the forest canopy, weakening or killing shade-intolerant species, particularly pines. Once kudzu gains access to the forest canopy, the liana formed can spread faster and more aggressively through a forest.


An extreme example of kudzu overgrowth of natural vegetation. Kerry Britton, USDA Forest Service,


Economic Impacts:

By outcompeting, smothering, and physically removing native vegetation, kudzu damages to lost forest production for southern commercial timber producers has been estimated to be as high as $48 per acre ($118 per hectare) per year. Kudzu control costs can be as high as $200 per acre per year. Control costs on power company rights-of-way and transmission equipment have been estimated as high as $1.5 million per year. Kudzu can also be a problem along highway rights-of-way.


Kudzu overgrowth of a southern highway embankment. Chris Evans, Illinois Wildlife Action Plan,



With a growth rate of up to one foot (0.3 meter) per day, simply controlling or managing kudzu can become a “fool’s errand” of never ending activity. In areas where the plant cannot be tolerated at all, kudzu control is basically kudzu eradication. To prevent reinvasion, complete eradication is required, which means every root crown on a site must be killed. Due to the numerous root crowns at vine nodes, eradication of a well-established population of kudzu could take 5 – 10 years of concentrated effort. The more mature the population, the more difficult eradication becomes as a result the numerous crowns and the large rhizome system that can store significant amounts of starch to feed the plant. Lianas are also more efficient at producing starch and sending it to the root system than are horizontal, ground-based vines.

Eradication of kudzu with herbicides calls for frequent defoliation during the growing season, while most of the plant’s energy is devoted to vine production and growth. Defoliation forces the plant to call on root starch reserves to resume foliage growth activities, helping to diminish reserves of starch and prevent storage of new reserves. If a single treatment is all that can be undertaken in a year, it should be implemented in early-fall as foliage starch allocation to the root system replenishing that used for growth during the spring and summer takes place in the early-fall.

If physical or mechanical control methods are selected, eradication of well-established kudzu populations could take many years or be ineffective in the long-term. Mechanical harvesting of kudzu foliage limits the production of new food reserves by reducing photosynthesis; regrowth helps to deplete starch stored in the root system. Mowing of trailing vines and root crowns every two weeks may take up to ten years to eradicate small, immature patches of kudzu, assuming that all root heads are mowed. Mowing is more likely to result in eradication if used with herbicide application. During mechanical eradication efforts, all cut plant material should be destroyed by burning or by bagging and landfilling.

The use of intensive conservation grazing by herbivores such as sheep or goats can help control young, tender kudzu growth and make control by herbicides more effective over shorter periods of time by helping to reduce energy reserves.

For information regarding appropriate use of herbicides against kudzu and other invasive plants, please consult The Nature Conservancy’s Weed Control Methods Handbook. Make certain to consult your state’s environmental conservation or natural resource management agency to determine which herbicides are legal for kudzu control in your state.


Herbicide spraying for kudzu control. James H. Miller, USDA Forest Service,



In the 1950s, the Agricultural Conservation Program removed kudzu from the list of species acceptable for use as an agricultural forage crop or soil stabilization plant. Congress listed kudzu as a Federal Noxious Weed in 1998. In 2014, the State of New York designated kudzu as a prohibited plant under the state’s Environmental Conservation Law.

Wild Parsnip

Introduction and Distribution  |  Habitat, Biology, and Ecology  |  Impacts  |  Prevention  |  Control and  Management


Wild parsnip (Pastinaca sativa). Leslie J. Mehrhoff, University of Connecticut,


Wild parsnip (Pastinaca sativa) is a biennial/perennial herb native to Eurasia. In appearance, it looks and smells quite like cultivated parsnip (in point of fact, wild parsnip is part of the Apiaceae (or Umbelliferae) family which includes carrots, celery, parsley, parsnip, Angelica, and Queen Anne’s Lace, most of which are aromatic plants with hollow stems). It is believed to be an escapee from parsnip that was originally under cultivation. The plant typically can grow up to 4 feet (1.2 m) tall in an average year. Wild parsnip is common throughout the northern United States and southern Canada. Its range reaches from Vermont to California and south to Louisiana (it is not found in Hawaii, Mississippi, Alabama, Georgia, and Florida). Reported populations can be found across New York State with the heaviest concentrations being found in the Lower Hudson Valley, Catskills, and southern Adirondacks.


North American range of wild parsnip (Pastinaca sativa). USDA PLANTS database, July 2014.

NYS range of wild parsnip. accessed 16 July 2014


Wild parsnip is an herbaceous plant which can grow from 4 – 5 feet (123 – 150 cm) tall. It can survive in a broad range of environmental settings, from dry soils to wet meadows. It grows best in rich, calcareous, alkaline, moist soils. It is commonly found growing along roadsides, in pastures, and in abandoned fields, or any place where the soil has been disturbed and native vegetation has yet to become fully established.

The roots are generally smooth and cylindrical, although sometimes lateral roots will grow out from the central tap root.

Seedlings emerge from February through April, form rosettes in their first year, and grow vegetatively for one or more years, at which time they will form an aerial shoot (called a “bolt”) and flower. Wild parsnip produces a rosette of broad, hairless, ovate, compound pinnate leaves, up to 6 inches (15.2 cm) in length, terminating with several pairs of leaflets with saw-toothed margins; they can grow up to 16 inches (40 cm) long. Leaflets are arranged in pairs along the stalk. Lower leaves have short stems, upper leaves are stemless. The leaves give off a pungent odor when crushed. During the vegetative growth season, wild parsnip continuously produces and loses leaves. The flower stalk develops from the rosette in the second year and can grow to a height of 4 – 5 feet (123 – 150 cm). It is grooved, hairy, and, except at the nodes, hollow. The stalk is sparsely branched. Over the winter above ground wild parsnip plants die back with only one or two leaves remaining on each plant.


First-year wild parsnip growth. Patrick J. Alexander @ USDA-NRCS PLANTS Database


Close-up of wild parsnip leaves. Patrick J. Alexander @ USDA-NRCS PLANTS Database


Wild parsnip stems are hollow except at the nodes. Missouri Dept. of Conservation (left);
Leslie J. Mehrhoff, University of Connecticut, (right)


Each wild parsnip plant produces hundreds of small yellow flowers which bloom from June to mid-July. The flowers are arranged in a loose compound umbel (a structure made up of a number of short flower stalks which spread from a common point, looking like the ribs of an umbrella). An umbel can measure from 4 – 8 inches (10 – 20 cm) in diameter. The flowers consist of five yellow petals curled inward, five stamens, and one pistil. The large, straw to light-brown seeds that are produced by the flower heads are round to oval, flat, slightly ribbed with narrow wings and are 1 ½ – 3 inches (4 – 8 mm) long. Seeds mature by early July. Plants die after producing seeds; the dead stalk will remain standing through the winter. Seeds can remain viable in the soil for four years. Seedling mortality is high; less than 1% of seedlings survive to mature and reproduce.



Wild parsnip second-year growth. Virginia Tech Weed ID Guide (left). Mature flowering wild parsnip. University of Massachusetts Extension (right)


Wild parsnip umbel. Virginia Tech Weed ID Guide

Wild parsnip seeds. Leslie J. Mehrhoff, University of Connecticut,


Click the table (above) for a 2-page ID guide to giant hogweed, native cow parsnip, native purple-stemmed Angelica, poison hemlock, and wild parsnip


Ecological Impacts:

Wild parsnip invades and modifies disturbed open habitats. Well-established fields and meadows are not likely to be invaded, but parsnip can become well-established along the edges and in disturbed areas. Once an infestation begins, it can spread into adjacent areas and form dense stands in high-quality fields and meadows. Wild parsnip is also very persistent on sites that remain disturbed or bare such as paths, roadsides, and utility rights-of-way.

Human Health Impacts:

While wild Parsnip roots are edible, the plant produces a compound in its leaves, stems, flowers, and fruits that causes intense, localized burning, rash, severe blistering, and discoloration on contact with the skin on sunny days. This condition, known as phytophotodermatitis, is caused by furanocoumarin contained in the sap. This is not an allergic reaction, it is a chemical burn brought on by an increase in the skin’s sensitivity to sunlight. Affected areas can remain discolored and sensitive to sunlight for up to two years, similar to but not as severe as contact with giant hogweed. This reaction is not brought on by contact with the foliage of the plant, only by contact with the sap.

Contact may occur when working, hiking, and harvesting crops, including when visiting u-pick operations. To reduce the risk of exposure to wild parsnip sap, when undertaking such pursuits one should wear long-sleeved shirts, gloves and long pants.

If one should come in contact with wild parsnip sap, you should immediately cover the exposed skin to prevent the reaction to sunlight (but the area will remain sensitized for about eight hours). The contact area should be washed with warm water and a mild soap. If exposure to sunlight causes a burn and blisters to develop the affected area should be covered with a cool, damp cloth to help relieve pain. The blistered skin should be kept out of the sunlight to avoid further burning. If blistering is severe, see a physician. There is no cure for parsnip burns; however, a topical or systemic cortisone steroid may relieve discomfort.


Wild parsnip burns. Andrew Link, Lacrosse Tribune 2013

The essential oil of parsnip roots contains a large percentage of Myristicine, a strong human hallucinogen.

Wild parsnip is regulated in Ohio, Illinois, Tennessee, and Wisconsin.


Remove new infestations while they are still small. Avoid mowing areas with wild parsnip when viable seeds are present as equipment readily spreads seed to new areas. Clean mowing equipment before moving from an area with wild parsnip to one without. When possible, plan to harvest/mow areas without wild parsnip before moving to fields where it is present. Time control efforts to prevent spread of the plant.


Management decisions should be based on the quality of the area, the degree of the infestation, and use of the infested area by people or livestock.

Manual control for small patches is effective. Cut the root 1” below the ground using a tool such as a spaded shovel or remove plants by hand pulling, gripping the stalk just above the ground. These control measures should be undertaken before wild parsnip plants go to seed. If hand pulling after seed formation, take steps to destroy the seeds. For small areas which have set seed, cut the tops with clippers, bag the seed heads in clear plastic and allow to rot.

Mowing – Mow when plants first produce flowers, but before seeds enlarge. At this stage plants have depleted their root resources and often die when cut. Some plants will re-sprout, so a follow-up mowing may be needed. When using any type of mowing equipment, take precautions to prevent plant sap from contacting exposed skin. Mowing can tend to favor wild parsnip rosettes as more sunlight is able to reach them, as well as reducing the number of plants competing with them for light and nutrients.

Chemicals – General-use herbicides such as glyphosate or triclopyr can be applied as spot treatments to basal rosettes. Be sure to follow all label and state requirements.

Biocontrol – No effective options are currently known. The parsnip webworm infests individual plants, but is not known to significantly damage large patches.

Plan to monitor the area long-term for seedlings emerging from the seed bank.

Whatever type of control method is employed, make certain to take measures to protect skin and eyes from contact with the plant’s sap.


Background  |  Origin and Expansion  |  Biology  |  Characteristics and Identification  |  Impacts  |  Prevention, Control, and Management



Mugwort  (Artemesia vulgaris) is an invasive perennial forb that is widespread throughout North America, though it is most common in the eastern United States and Canada. It is a weed of nurseries, turfgrass, vineyards, waste areas, forest edges, and roadsides. Mugwort spreads aggressively through an extensive rhizome system and will readily form large, mono-specific stands.

Origin and Expansion

Mugwort is native to Europe and eastern Asia, where it has historically been used as a medicinal herb. Seed may have been first introduced to North America as early as the 16th century by Jesuit missionaries in Canada. It was also introduced throughout the continent as a contaminant in ship ballast and nursery stock.


Mugwort is a perennial with an extensive rhizome system. Shoots emerge during the spring, and flowering occurs from July to late September. A single plant can, depending on its environment, produce up to 200,000 seeds.  The small seeds (~1mm in diameter) are largely wind dispersed. Seed production does not seem to be a major factor in the spread of mugwort populations, however, and some biotypes do not produce viable seed.  Instead, mugwort spreads largely through vegetative expansion and the anthropogenic dispersal of root propagules. The root system is extensive though shallow (to 20 cm in depth), with numerous branching rhizomes up to 1 cm in diameter. Plants can regenerate from rhizome fragments as small as 2 cm (Klingeman et al. 2004).

Characteristics and Identification

The rarely-seen seedlings have oblong cotyledons without petioles. Adult stems are smooth and longitudinally ridged, with numerous axillary branches towards the upper portions of the plant. The stems become somewhat woody as they age. The leaves are alternate, densely covered with wooly, silver-white hairs on the underside, and slightly hairy on the upper surface. Leaf morphology is variable throughout the plant. The lower leaves are petiolate, with stipules at the base, and generally coarsely toothed and pinnately lobed. The upper leaves are sessile and lanceolate with smooth or toothed margins. The numerous ray and disk flowers are small (5 mm), green, and grow in racemes and clusters at the end of stems and branches. The foliage is aromatic and slightly pungent.


Mugwort is a problematic weed in nurseries, where small root fragments can easily contaminate nursery stock. It is also a major weed in turf grass, field-grown ornamental crops, and orchards. Stands of mugwort displace native species, and can delay or disrupt succession in natural ecosystems (Barney and DiTommaso 2003). Mugwort produces several terpenoid potential allellochemicals, and decaying mugwort foliage has been shown to inhibit the growth of red clover in laboratory experiments (Inderjit and Foy 1999). Mugwort pollen is a common cause of hay fever.

Prevention, Control, and Management

The dense root system of mugwort can make it difficult to control. Pulling is ineffective, and may even promote growth by leaving residual rhizome fragments in the soil. Mugwort tolerates mowing, and even sustained mowing over two years will not fully eradicate mugwort stands. The relatively shallow roots make mugwort vulnerable to repeated cultivation in agricultural systems, though this practice risks spreading root propagules.

Chemical control of mugwort can have limited effectiveness. Though non-specific broadleaf herbicides such as glyphosate or dicamba can effectively control mugwort, the rates required for adequate suppression are rarely economical (Bradley and Hagood 2002). For small infestations, multiple spot-treatments of glyphosate can be effective (Bing 1983).