Commonly grown and transplanted for its showy yellow flowers, yellow flag iris (Iris pseudacorus) 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, http://www.eddmaps.org/; accessed September 29, 2015).
Biology
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.
Description
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, Bugwood.org).Yellow Flag Iris seeds (Photo: Leslie J. Mehrhoff, University of Connecticut, Bugwood.org)
Impacts
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, Bugwood.org)
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.]
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
The Eurasian forb purple loosestrife, Lythrum salicaria, is an erect, branching, perennial that has invaded temperate wetlands throughout North America. It grows in many habitats with wet soils, including marshes, pond and lakesides, along stream and river banks, and in ditches. Purple loosestrife is also capable of establishing in drier soils, and may spread to meadows and even pastured land. It prefers full sun, but can grow in partially shaded environments. Purple loosestrife stem tissue develops air spaces between cells, allowing them to respire when partially submerged in water.
Stand of mature purple loosestrife. [Photo: Lesley J. Mehrhoff, University of CT, Invasive.org]
Origin and Expansion
Purple loosestrife is native to Europe, Asia and northern Africa, with a range that extends from Britain to Japan. Purple loosestrife was probably introduced multiple times to North America, both as a contaminant in ship ballast and as an herbal remedy for dysentery, diarrhea, and other digestive ailments. It was well-established in New England by the 1830s, and spread along canals and other waterways. Purportedly sterile cultivars, with many flower colors, are still sold by nurseries. Its range now extends throughout Canada and to all states but Hawaii and Florida.
North American range (December 2015). Map: USDA NRCS PLANTS Database
Biology
Purple loosestrife is a perennial, with a dense, woody rootstock that can produce dozens of stems. Shoot emergence and seed germination occurs as early as late April, and flowering begins by mid-June. Seedlings grow rapidly, and first year plants can reach nearly a meter in height and may even produce flowers. The flowers are insect-pollinated, principally by nectar feeders like bees and butterflies. Seed development begins by late July and continues throughout the season and into autumn. A single plant can produce over 2 million seeds. Senescence occurs with the first frost, and dead stems persist throughout the winter.
Though purple loosestrife seeds may not be particularly long-lived in the seed bank (they can survive for at least 3 years), the sheer number of seeds produced allows them to readily capitalize on disturbance. Seeds are dispersed by wind for short distances, by floating, and by anthropogenic means. Germination is best in wet, open soil under relatively warm temperatures (greater than 68°F). The seeds are capable of germinating and establishing under standing water. Though the rootstock buds prolifically, purple loosestrife does not generally spread through vegetative reproduction. Stem fragments can regrow, however, and mowing or otherwise damaging the plants may spread vegetative propagules.
Purple loosestrife flowers in full bloom. [Photo: Lesley J. Mehrhoff, University of CT, Invasive.org]Purple loosestrife flowers in full bloom. [Photo: Lesley J. Mehrhoff, University of CT, Invasive.org]
Description
Seedlings have oval cotyledons with long petioles. The stalkless stem leaves are 5-14 cm long, lance-shaped, and opposite. Leaf pairs often grow at 90 degree angles from one another, and leaves near the flowers are sometimes alternate. Stems are upright, angular, and densely hairy. Mature plants can reach up to 4m in height, and older plants often appear bush-like, with sometimes dozens of woody stems growing from a single rootstock. The showy purple flowers have 5-7 petals and grow in pairs or clusters on 10-40 cm tall spikes. Seeds are small (less than 1 mm in length) and lack an endosperm.
Purple loosestrife stalk and leaves. Photo: Ohio State Weed Lab, The Ohio State University, Invasive.org.Purple loosestrife stalk and leaves. Photo: Theodore Webster, USDA Agricultural research Service, Invasive.org
Impacts
Purple loosestrife is competitive and can rapidly displace native species if allowed to establish. Once established, the prolific seed production and dense canopy of purple loosestrife suppresses growth and regeneration of native plant communities. Monotypic stands of purple loosestrife may inhibit nesting by native waterfowl and other birds. Other aquatic wildlife, such as amphibians and turtles, may be similarly affected. The dense roots and stems trap sediments, raising the water table and reducing open waterways, which in turn may diminish the value of managed wetlands and impede water flow.
Large stands of purple loosestrife can clog waterways and deprive native wildlife of habitat and food. [Photo: Lesley J. Mehrhoff, University of CT, Invasive.org]
Control
Small infestations can be pulled by hand, though care must be taken to completely remove the root crown. Glyphosate or triclopyr based herbicides can also effectively control small stands, but as they are expensive and non-selective they are generally unsuitable for large purple loosestrife infestations. Mechanical or chemical management will require multiple years to completely remove adult plants and exhaust the seedbank.
Four species beetles (2 leaf beetles and 2 weevils) have been released in the United States as biocontrol agents for purple loosestrife. They have had some measure of success controlling purple loosestrife populations. The leaf-feeding beetles Galerucella calmariensis and G. pusilla defoliate and attack apical buds as both adults and larvae and can slow growth and diminish seed production. The weevil Nanophyes marmoratus feeds on seeds and flower buds, and the weevil Hylobius transversovittatus attacks both roots (as larvae) and foliage (as adults).
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
Didymo (Didymosphenia geminata), a globally invasive single-celled algae (diatom), is threatening the streams and rivers of New York State. Didymo secretes massive amounts of branching stalks, creating dense mats that cover the bottoms of streams and rivers. Nicknamed “rock snot” for its gooey appearance, didymo has been confirmed at eight locations in New York State since 2007.
A didymo bloom observed in the Batten Kill. – VT DEC
Origin & Spread
Historically, didymo was considered a widely distributed, yet uncommon, algal species native to the cool, running freshwaters of the northern hemisphere, including northern parts of North America, Europe, and Asia. Records collected over the last 150 years indicate that didymo widely occurred across Europe, including the UK, Norway, and northwest Russia. Didymo diatoms have been reported in the western US for over 100 years, but not as nuisance blooms.
Yet, within the last two decades, didymo blooms have been reported with increasing frequency and intensity across the globe, particularly in locations where historical blooms have not been previously recorded. In North America, didymo blooms were first documented in the 1990s in rivers on Vancouver Island, British Columbia. In the last 20 years, bloom occurrences have spread east across North America and have become increasingly common in the northeastern US, particularly in streams and rivers frequented by anglers and other aquatic recreationists. In 2011, the species could be found in 18 US states and 3 Canadian provinces.
Density of Obeservations, Rock Snot. iMapInvasives.org.
Didymo has been confirmed in eight locations in New York State since 2007, including:
Batten Kill and one tributary (Washington County)
Kayderosserras Creek (Saratoga County)
East Branch Delaware River below Pepacton Reservoir (Delaware County)
West Branch Delaware River below Cannonsville Reservoir (Delaware County)
West Branch Delaware River below Delhi to Cannonsville Reservoir (Delaware County)
Mainstem Delaware River (Delaware and Sullivan Counties)
Mouth of Little Delaware River (Delaware County)
Esopus Creek downstream of the Shandaken Portal (Ulster County)
All of these confirmed sites are prime fishing access points where the species has been observed by numerous anglers; it is very possible that Didymo exists in other waterways where it has yet to be identified.
View Didymo in NYS in a larger map Confirmed Didymo sites in New York State, 2010.
Use the navigation tools in the upper left corner of the map to move the map up, down, left or right or to zoom in or out. To go to full-size Google map click here.
Didymo bloom observed in Esopus Creek near Mt. Tremper, NY. – David Richardson, SUNY New Paltz
Didymo was first detected in the southern hemisphere in New Zealand in 2004, and has continued to spread through numerous watersheds on the South Island (over 32). New Zealand’s didymo blooms have been particularly severe, with mats growing up to 8 inches thick and extending over 2.5 miles in length. Consequently, the government agency Biosecurity New Zealand has been a global leader in efforts to slow the spread and educate the public about didymo.
Didymo has rapidly spread through streams on New Zealand’s South Island. Didymo in stream on New Zealand’s South Island. – New Zealand Fish and Game
In 2010, the first didymo blooms in South America were confirmed in several rivers flowing through Patagonia (Chile and Argentina). It was first detected there in the Futaleufú River, a site popular with kayakers from all over the world.
Humans are largely responsible for the recent and prolific spread of didymo beyond its historical range. The microscopic diatoms can cling to fishing gear, waders, boots, and boats, and are capable of surviving at least 40 days outside of a stream as long as they remain in a damp, cool environment.
Felt-soled wading boots are major vectors of didymo as the soles absorb cells like a sponge and provide a temporary habitat for the didymo cells until an angler visits a new site. As a result, natural resource agencies as well as fishing organizations and supply stores like Trout Unlimited and L.L. Bean have promoted non-felt wading boot alternatives. Furthermore, the states of Vermont, Maryland and Alaska have outlawed the use of felt-soled wading boots.
Felt-soled wading boots (commonly used by fly fishers) are known vectors of didym0. Felt-soled wading boots – Treehugger.com
Biology
Didymo is a freshwater diatom, a type of single-celled algae that lives attached to rocks and other substrates on the bottom of streams. Under certain conditions, didymo will secrete thick, branching stalks (composed of polysaccharides and protein) outside its cells that form dense tangled mats. The resulting didymo blooms can completely dominate streambeds, capable of lasting several months, with mats greater than 8 in thick and over 0.5 mi long. Unfortunately, the triggers of excessive stalk formation – possibly genetic and/or environmental – are unknown and the subject of current scientific research.
Didymo is a large freshwater diatom that can secrete thick, branching stalks. Close up of Didymo stalks – Sarah Spaulding, USGS
Thick didymo mats resemble fiber-glass insulation or wet toilet paper, inspiring its nickname, “rock snot.” It is generally light tan to brown in color (not green), with stalks sometimes forming long white strands. Clumps of didymo are not slimy, resemble wet wool, and are tough to pull apart.
Didymo blooms exhibit a dense woolly appearance, with stalks often forming long white tails that resemble wet toilet paper. South Dakota Dept. of Environment & Natural Resources
In contrast to other types of freshwater algae that form nuisance blooms, didymo blooms first appeared in streams with relatively high water quality (clear, cold, low nutrients, stable flow). Often blooms occurred immediately downstream of impoundments, where flows were generally stable, regulated, and nutrient poor. More recently, as its range has expanded, didymo has been recorded in warmer, more nutrient-enriched waters, not associated with dams.
Impacts
Didymo can alter the diversity and distribution of native stream species and may have negative consequences on how stream ecosystems function.
The extensive stalks produced by didymo cells persist in invaded streams longer than the diatoms themselves and are resistant to degradation; reports from Colorado indicate thick mats of didymo stalks can persist up to 2 months after peak growth. These mats may trap stream sediments, changing the physical nature of the stream bottom and affecting the ability of native stream algae to colonize.
By covering and dominating the substrate, didymo may alter habitat and available food resources for bottom-dwelling stream invertebrates. Recent studies in New Zealand and North America have reported shifts in the invertebrate community as a result of didymo invasion. The proportion of larger sized, environmentally sensitive insect groups like mayflies, stoneflies, and caddisflies (Ephemeroptera, Plecoptera, Trichoptera; known collectively as EPT taxa) tend to decline in didymo infested waters while the proportion of smaller invertebrates, particularly midge larvae (Diptera: Chironomidae), worms (Oligochaeta, Nematoda), and crustaceans (Cladocera), increases. In most cases, the total number of invertebrates actually increases with didymo invasion, possibly as a result of the new habitat or additional food resources provided by the dense didymo stalks.
Shifts in the abundance and diversity of invertebrates as a result of didymo invasion could potentially affect the fish that feed on them. Declines in mayflies, stoneflies and caddisflies could threaten trout populations. Furthermore, didymo blooms covering large stretches of the stream bottom could alter stream habitat that is important for fish feeding and spawning. Research on the effects of didymo on native and sport fisheries is still underway. To date, research in North America and New Zealand has been inconclusive, with some studies suggesting no impact of didymo on fish growth and productivity, while others have observed native fish population decline in didymo-invaded waters.
Didymo invasions, although unsightly, do not produce an odor or threaten human health. Infestations do have significant negative impacts on all water-associated recreational activities, particularly sport-fishing. Floating didymo stalks tangle up lines, flies and lures. Additionally, didymo blooms have blocked water intake pipes and canals. Consequently, didymo remains a serious economic concern for fisheries, tourism, irrigation, and hydropower.
Detection
Didymo can exist both as a single cell without stalks or as a colony of cells with branching, mat-forming stalks. Unfortunately, invasions are not usually detected until dense didymo mats occur. Researchers who have intensively sampled stream algae by scraping rocks and filtering water from streams with no previous history of didymo blooms have identified single didymo cells when examining those samples under the microscope. Yet, this type of work is extremely time and labor-intensive as didymo cells can be quite rare. Research is now ongoing to develop DNA-based tests for detecting didymo in streams, even at very low concentrations.
Prevention and Management
Currently, there are no methods available for controlling or eradicating didymo once it has infested a water body. Research in New Zealand is underway to identify and evaluate the safety and effectiveness several didymo-killing chemicals.
Spread prevention is, therefore, the only method we have for protecting our streams and rivers from didymo invasion. Water recreationists must take great care to inspect, clean, and dry all equipment, especially waders and boots when leaving an infested stream or river.
Inspect: Look for and remove all clumps of algae and discard in designated invasive species disposal stations or upland.
Clean: Clean and disinfect any equipment that has come in contact with the water, whether you observe a didymo bloom or not as other aquatic invasive species and diseases may hitchhike on your equipment. Waders and gear should be soaked and/or scrubbed with a 2-percent solution of household bleach or a 5-percent solution of detergent and very hot tap water (> 115°F); ‘eco-friendly’ detergents are not recommended. Absorbent materials (e.g., felt soles, foam) will require prolonged soaking to kill didymo cells (> 30 minutes). Gear could also be placed in a freezer until all of the moisture is frozen solid (note: freezing may damage some gear and will only kill didymo, not necessarily invasive fish diseases).
Dry: Drying will kill didymo, but this method alone is not recommended for absorbent materials because didymo can survive in slightly moist environments for an extended period of time.
If cleaning, fully drying, or freezing is not practical, restrict equipment use to a single water body. One didymo cell transported on gear could result in an invasive didymo bloom – thus, precautionary action is essential!
Photo and Map Credits
Didymo bloom observed in the Batten Kill. – VT DEC
North American distribution of didymo 21 July 2008. National Invasive Species Information Center. Modified from map created by Karl Hermann, Sarah Spaulding, and Tera Keller. For original image click here
Didymo bloom observed in Esopus Creek near Mt. Tremper, NY. – David Richardson, SUNY New Paltz
Didymo in stream on New Zealand’s South Island. – New Zealand Fish and Game
Felt-soled wading boots – Treehugger.com
Close up of Didymo stalks – Sarah Spaulding, USGS
Didymo blooms showing dense woolly appearance – South Dakota Dept. of Environment & Natural Resources
The non-native Phragmites australis, or common reed, can rapidly form dense stands of stems which crowd out or shade native vegetation in inland and estuary wetland areas. Phragmites turns rich habitats into monocultures devoid of the diversity needed to support a thriving ecosystem. Non-native Phragmites can alter habitats by changing marsh hydrology; decreasing salinity in brackish wetlands; changing local topography; increasing fire potential; and outcompeting plants, both above and belowground. These habitat changes threaten the wildlife that depend on those wetland areas for survival.
Stand of Phragmites Credit: Bernd Blossey, Cornell University, Bugwood.org
History
Common reed, Phragmites australis, is in the Poaceae or grass family. There are at least three lineages, or strains, of common reed in the U.S. At least one is native to the U.S. including the one that was most common in New York, P. australis subsp. americanus. Another common reed strain, P. australis var. berlandieri may or may not be native to the U.S. and is found in California, along the Gulf Coast, and in the Southeast. One strain is non-native, and was accidentally introduced from Europe in the late 18th or early 19th century in ship ballast. This non-native strain is now the most common Phragmites found in New York and the Northeast. There is no field evidence that the non-native will hybridize with the native Phragmites at this time. This fact sheet focuses on the non-native Phragmites.
Biology
The non-native Phragmites is a perennial grass that can reach over 15 feet in height. It is often found in dense clonal stands made up of living stems and standing dead stems. Stems of the non-native Phragmites are hollow, usually green with yellow nodes during the growing season, and yellow when dry in the winter. Phragmites leaves are blue-green to yellow-green, up to 20 inches long and 1 to 1.5 inches wide at their widest point. They are arranged all along one side of a stem.
In late July and August, Phragmites is in bloom with purple to gold highly-branched panicles of flowers. The seeds are grayish and appear fluffy due to the silky hairs that cover each seed. Spread occurs through rhizomes, stolons, and seeds; stolons can grow up to 43 feet from the parent plant.
Phragmites in bloom Credit: Leslie J. Mehrhoff, University of Connecticut, Bugwood.org
Root growth belowground is also profuse. Phragmites forms a ticket of roots and rhizomes that can spread 10 or more feet and several feet deep in one growing season.
Each Phragmites plant produces thousands of seeds each year, but seed viability is low, although viability varies from year to year. New sites are established through seed movement and from rhizome fragments that float down stream or are moved in soil, especially along roadsides.
Large clumps of Phragmites can live for decades, but no part lives for more than 8 years.
There are physiological differences between the native Phragmites and the non-native Phragmites. See the Plant Conservation Alliance Phragmites Fact Sheet comparison table for details.
Phragmites in seed. Credit: Jil Swearingen, USDI National Park Service, Bugwood.org
Habitat
The non-native Phragmites occurs throughout the eastern half of the U.S. and in Colorado. In New York, Phragmites is ubiquitous, growing in roadside ditches and swales; tidal and non-tidal wetlands; freshwater and brackish marshes; river, lake and pond edges; and disturbed areas. It tolerates fresh and moderately-saline water and prefers full sun.
Credit: John M. Randall, The Nature Conservancy, Bugwood.org
Credit: John M. Randall, The Nature Conservancy, Bugwood.org
Management
Due to the similarity of non-native Phragmites and native Phragmites, proper identification of the grass is important before taking management action. Due to Phragmites growth in sensitive habitats, be sure to have a restoration plan in place for the area once Phragmites has been eliminated. Phragmites roots hold onto soil, and clonal colonies trap nutrients and organic matter and add to the organic matter in the soil. After Phragmites colonies are removed the soil may be more prone to erosion.
To control Phragmites a number of tactics may be used, but due to the many variables at each site many suggest that Phragmites management should be “site-specific, goal-specific, and value-driven.” Often multiple tactics are needed to ensure success. The best time to manage Phragmites is in midsummer when it is releasing pollen. Thorough monitoring and follow-up management are necessary to control shoots from surviving rhizomes.
Prevention
Maintain, or plant, vegetation that competes with Phragmites. Jesuit’s bark (Iva frutescens), groundsel-tree (Baccharis halimifolia), black rush (Juncus roemerianus), and saltmeadow cordgrass (Spartina patens) have been shown to limit Phragmites spread. Also, reducing nutrient loads may restrict the spread of Phragmites.
Mechanical
Repeated mowing may produce short-term results and repeated stem breakage in high-water years has been shown to kill large portions of Phragmites colonies. Hand pulling is not feasible due to the expansive and tough root and rhizome network. Root removal from the soil is not effective as small or broken portions of rhizomes left in the soil can create new plants.
Hydrologic
Manipulating the water level around Phragmites has been shown to decrease populations in some conditions. Consult the Element Stewardship Abstract for Phragmites australis produced by the Nature Conservancy for more information. http://www.invasive.org/gist/esadocs/documnts/phraaus.pdf
Chemical
There are herbicides available for Phragmites control. New colonies, with smaller root and rhizome systems, are easier to control with herbicides. Apply after the plant has flowered, in late summer or early fall. Applications can be foliar, cut stump, or injected. Multiple years of treatment may be necessary to eliminate any surviving rhizomes. Herbicides applied in wetland areas must be applied by a certified pesticide applicator. Contact your local Cornell Cooperative Extension office, http://www.cce.cornell.edu, for herbicide usage assistance. Always apply pesticides according to the label directions; it’s the law.
Fire
Prescribed burns have been shown effective when conditions are right, and can occur in conjunction with herbicides or water level management. To be successful as a stand-alone tool, burns need to be hot enough to kill rhizomes in the soil. After herbicide treatments, burns can remove standing dead stems to make way for desirable vegetation. Flooding after burns will limit soil air to surviving rhizomes. Burns should be conducted once flowering has occurred. For more information on controlled burns, see the USDA Forest Service Fire Effects Information System “Phragmites australis Fact Sheet,” Fire Effects section at http://www.fs.fed.us/database/feis/plants/graminoid/phraus/all.html#FIRE%20EFFECTS.
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
A Patch of floating water chestnut (Trapa natans) leaves.
Background
If a shoreline property owner in New York or the Northeast complains to you about their water chestnut problem, don’t think they are talking about Chinese takeout. The European water chestnut (Trapa natans), an invasive aquatic plant released inadvertently into waters of the Northeast in the late 1800s, is slowly but inexorably spreading throughout New York State, clogging waterways, lakes and ponds and altering aquatic habitats.
It must be pointed out that this plant species is not the same as the water chestnut which can be purchased in cans at the supermarket. The fruits of T. natans, however, are used as a source of food in Asia and have been utilized for their medicinal (and claimed) magical properties.
T. natans is native to Europe, Asia and Africa. In its native habitat, the plant is kept in check by native insect parasites. These insects are not present in North America and the plant, once released into the wild, is free to reproduce rapidly. T. natans colonizes areas of freshwater lakes and ponds and slow-moving streams and rivers where it forms dense mats of floating vegetation, causing problems for boaters and swimmers and negatively impacting aquatic ecosystem functioning.
Common names: horned water chestnut, water caltrop
Introduction History and Distribution
T. natans is native to Western Europe and Africa and northeast Asia, including eastern Russia, China, and southeast Asia to Indonesia. T. natans was first introduced to North America in the mid- to late-1870s, when it is known to have been introduced into the Cambridge botanical garden at Harvard University around 1877. It is known to have been planted in other ponds in that area, as well, and also in Concord, MA, in a pond near the Sudbury River. The plant escaped cultivation and was found growing in the Charles River by 1879. The plant was introduced into Collins Lake near Scotia, NY (in the Hudson River-Mohawk River drainage basin) around 1884, possibly as an intentional introduction for waterfowl food or possibly as a water garden escapee.
By the early part of the 1900s, water chestnut was established in the Hudson River. The first Great Lakes Basin introductions were sometime before the late-1950s when the plant was discovered growing in Keuka Lake (one of NY’s Finger Lakes). A major infestation of more than 300 acres exists throughout some 55 miles of Lake Champlain between New York and Vermont. Water chestnut can now be found throughout NY, from the Niagara Frontier through the Finger Lakes, from Lake Champlain to Long Island.
The North American distribution of water chestnut now extends throughout New England, south as far as Virginia, California, and in the Canadian Province of Quebec in a tributary of the Richelieu River. The plant has the potential to spread into the warmer regions of the U.S. as far south as Florida.
North American Distribution of water chestnut as of September 2014.
Identification and Biology
T. natans is a rooted aquatic annual herb that dies back at the end of each growing season. Re-growth is by means of seeds that germinate in the spring. Each seed produces 10 to 15 stems with submerged and floating leaves, terminating in floating rosettes. The feathery submersed leaves can be up to six inches (15 cm) long, and are alternate on the stem forming whorls around the stem. The three-quarter to one and a half inch (2 – 4 cm) glossy green floating leaves are triangular with toothed edges and form rosettes around the end of the stem. The floating leaves also have prominent veins and short, stiff hairs on their lower surface. The petioles (the stalks attaching the leaf blade to the stem; the transition between the stem and the leaf blade) of the floating leaves are two to eight feet (0.6 – 2.4 m) and contain spongy, buoyant bladders, allowing the rosettes to float on the surface of the water. Stems can reach lengths of up to 16 feet (4.9 m), although typical lengths tend to be in the six to eight foot (1.8 – 2.4 m) range. The stems are anchored to the bed of the waterbody by numerous branched roots. Single small, white flowers with four one-third inch (8.3 mm) long petals sprout in the center of the rosette.
Trapa rosette showing nuts and inflated leaf petioles
Each rosette is capable of producing up to 20 hard, nut-like fruits. Water chestnut starts to produce fruits in July; the fruits, which ripen in about a month, each contain a single seed. The 1 to 1.5 inch (2.5 – 4 cm) wide fruits grow under water and have four very sharp spines. Water chestnut seeds generally fall almost directly beneath their parent plants and serve to propagate the parent colony. Population overwintering is accomplished through mature, greenish brown nuts sinking to the bottom where they can remain viable in the sediment for up to 12 years. Some seeds, however, or plant parts (floating rosettes) that still contain nuts, may be moved downstream in currents. Ducks, geese and other waterfowl may also play a role in the nuts’ dispersal (the spiny nuts have been observed tangled in the feathers of Canada geese). Old nuts, black in color, will float, and are not viable. When deposited in shallow water or on the shore, water chestnut nuts can lead to injuries if stepped on.
Ecological Impacts
Water chestnut has become a significant environmental nuisance throughout much of its range, particularly in the Hudson, Connecticut and Potomac Rivers, and in Lake Champlain. The plant can form nearly impenetrable floating mats of vegetation. These mats create a hazard for boaters and other water recreators. The density of the mats can severely limit light penetration into the water and reduce or eliminate the growth of native aquatic plants beneath the canopy. The reduced plant growth combined with the decomposition of the water chestnut plants which die back each year can result in reduced levels of dissolved oxygen in the water, impact other aquatic organisms, and potentially lead to fish kills. The rapid and abundant growth of water chestnut can also out-compete both submerged and emergent native aquatic vegetation.
Water chestnut infestation on Lake ChamplainA massive riverine infestation of water chestnut.
Water chestnut has little nutritional or habitat value to fish or waterfowl and can have a significant impact on the use of an infested area by native species.
T. natans likely impacts non-native and invasive plant and animal species in the same manner it impacts natives. Some of the potentially impacted invasive plant species might include: Eurasian watermilfoil (Myriophyllum spicatum), curly pondweed (Potamogeton crispus), and Eurasian or brittle water-nymph (Najas minor). It is not yet known in a match up of T. natans or and hydrilla (Hydrilla verticillata, which invader would outcompete which. Because of its invasiveness and severity of its impacts, T. natans has been listed in federal regulations prohibiting interstate sale/transportation of noxious plants.
Economic Impacts
Economic impacts result from T. natan’s impenetrable mats of vegetation which can impede swimming, boating, commercial navigation, fishing, and waterfowl hunting. Untreated populations of such an aquatic invasive species also can result in losses to shoreline property values and, as a result, to local government property tax revenues. As mentioned earlier, the sharp, spiny nuts can result in puncture injuries to swimmers and recreators walking along the shore of infested areas and can injure the feet of livestock and horses, as well.
One example of the cost of managing T. natans in a waterbody is the experience of the States of New York and Vermont on Lake Champlain. From 1982 through 2011, $9,600,000 has been spent on Trapa control in the lake with funding from a number of sources including: the two states; the U.S. Army Corps of Engineers; the U.S. Fish and Wildlife Service; the U.S. Department of Agriculture; Ducks Unlimited; the Lake Champlain Basin Program; and The Nature Conservancy. A combination of hand pulling and mechanical harvesting has been used on the lake since the early-1980s. Significant reductions of T. natans populations resulted from this prolonged annual control effort, however, every time that funds were reduced, rapid grow back of the species and extension of its range in the lake was observed.
Lake Champlain, NY, water chestnut relative annual control costs, 1982 – 2011
Control
Mechanical and Chemical Control
Mechanical Harvester
It is much easier and less expensive to control newly introduced populations of T. natans. Early detection of introductions and a rapid control response are key to preventing high-impact infestations. Because T. natans is an annual plant, effective control can be achieved if seed formation is prevented. Small populations can be controlled by hand pulling working from canoes or kayaks.
Large infestations usually require the use of mechanical harvesters or the application of aquatic herbicides. Regardless of treatment type, it should ideally take place before the fruit has ripened and dropped to the bottom forming a long-term seed bank. Because of the potential of unintentional spread of floating plant parts offsite, mechanical harvesting should be undertaken only by trained and certified equipment operators. Since water chestnut overwinters entirely by seeds that may remain viable in the sediment for up to 12 years, repeated annual control is critical to deplete the seed bank. Treatment generally is needed for five to twelve years to ensure complete eradication and can be very expensive (see Economic Impact, above).
Potential negative impacts to non-target species and public perceptions regarding the use of chemicals in recreational waters have limited chemical control of T. natans except as a treatment of last resort and usually only in still or sluggishly flowing waters. The herbicide 2,4-D has been tested and shown to be non-adverse on non-target species. 2,4-D has been used widely in the U.S. Another herbicide that is effective on T. natans is glyphosate. Application of aquatic herbicides requires both a licensed pesticide applicator and a permit from your state environmental regulatory agency.
Biological Control
The unfortunate fact is that for large infestations of water chestnut (i.e. those too large to be controlled by hand-pulling) over the long-term mechanical and chemical control measures have proven to be impractical to provide an economically sustainable control of water chestnut. Scientists have now turned to the potential of biocontrol agents to serve as a long-term solution to water chestnut infestations.
A number of potential biological control agents were found in field surveys in the native European and Asian ranges of water chestnut. The most promising biocontrol species appeared to be the leaf beetle Galerucella birmanica. Unfortunately, field observations in China suggested that G. birmanica may also attack native water shield (Brasenia schreberi) in addition to Trapa natans. This host non-specificity could be problematic to the use of the beetle for biocontrol in North America.
Laboratory and field tests initially indicated that out of 19 different plant species in 13 different families, G. birmanica laid eggs and completed development only on species of Trapa and B. schreberi. Adult G. birmanica in the field and lab indicated that the beetles showed a strong preference for T. natans. This preference continued even after the water chestnut was completely defoliated; adults resisted migrating to nearby water shield. While this is very promising news, additional studies on host specificity with additional North American aquatic plants are on-going. [Ding, et. al., 2006]
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
References
Blossey B, Schroeder D, Hight S, Malecki R. 1994. Host specificity and environmental impact of two leaf beetles (Galerucella calmariensis and G. pusilla) for biological control of water chestnut (Trapa natans). Weed Science 42:134-140.
Deck J, Nosko P. 2002. Population establishment, dispersal, and impact of Galerucella pusilla and G. calmariensis, introduced to control water chestnut in central Ontario. Biological Control 23: 228-236.
Ding J, Blossey B, Du Y, Zheng F. 2006. Galerucella birmanica (Coleoptera: Chrysomelidae), a promising potential biological control agent of water chestnut, Trapa natans. Biological Control. Vol.36, Issue 1, Pages 80–90
Fernald ML. 1950. Gray’s Manual of Botany. 8th ed. American Book Company, N.Y.
Gleason HA. 1957. The New Britton and Brown Illustrated Flora of the Northeastern U.S. and Adjacent Canada. New York Botanical Gardens, N.Y.
Methe BA, Soracco RJ, Madsen JD, Boylen CW. 1993. Seed production and growth of water chestnut as influenced by cutting. J. Aquat. Plant Manage. 31: 154-157.
Mills EL, Leach JH, Carlton JT, Secor CL. 1993. Exotic species in the Great Lakes: A history of biotic crises and anthropogenic introductions. Journal of Great Lakes Research 19: 1-54.
Mullin BH. 1998. The biology and management of water chestnut (Trapa natans L.). Weed Technology 12:397-401.
Pemberton RW. 2002. Water Chestnut. In: Van Driesche R., et al. 2002. Biological Control of Invasive Plants in the Eastern United States. USDA Forest Service Publication FHTET-2002-04.
Rawinski T. 1982. The ecology and management of water chestnut (Trapa natans L.) in central New York. M.S. thesis, Cornell University.
Vermont Invasive Exotic Plant Fact Sheet Series: Water Chestnut. Vermont Agency of Natural Resources and The Nature Conservancy, Vermont Chapter. June, 1998.
Hunt T, Marangelo P. 2012. 2011 Water Chestnut Management Program: Lake Champlain and Inland Vermont Waters, Final Report. Vermont Department of Environmental Conservation. May 2012.
Photo Credits
Patch of floating water chestnut (Trapa natans) leaves. John M. Randall, The Nature Conservancy, www.forestryimages.org
Eurasian watermilfoil, Myriophyllum spicatum L., (Fig. 1) is a submersed aquatic plant that has become a major aquatic invader across much of North America. Plants are rooted at the lake bottom and grow rapidly creating dense beds and canopies (Fig. 2). Milfoil typically grows in water 1 to 4 meters (3.2 to 13 feet) deep, but has been found in water as deep as 10 m (32.8 ft). Stem densities can exceed 300/m2 (359/yd2) in shallow water. Conventional control efforts such as mechanical harvesting have been unsuccessful in providing more than short-term relief. The use of herbicides have been found to suppress regrowth for six weeks to a year but have considerable negative effects on non-target organisms (e.g., mortality of native macrophytes, fish kills, increased algal growth, and contamination of public water supplies). Mechanical harvesting can result in short term localized population reductions but these methods are labor intensive and costly; continued relief must be supported by long-term mechanical intervention. Further, since harvesting inevitably results in the release of milfoil fragments, the harvesting process itself may be responsible for spreading the plant to uninfested areas.
Figure 2. Dense Myriophyllum spicatum canopy on Cayuga Lake, Ithaca, NY. (Photo: Robert L. Johnson, Bugwood.org)
Nature of Damage
Ecologic damage. Introduction of Eurasian watermilfoil can result in native macrophyte diversity and abundance declines. Eurasian watermilfoil beds form dense canopies at the water surface thereby reducing light penetration early in the season before native macrophytes have reached their full growth, shading them out and slowing/reducing growth potential. Eurasian watermilfoil beds, as a result of the reduction in native plants, have been found to contain significantly fewer macroinvertebrates (including benthic invertebrates) and a concomitantly lower abundance of native fish species. Milfoil-infested lakes tend to have reduced fish spawning areas and lowered fish growth rates. Native waterfowl in the Great Lakes have been found to avoid foraging for food in beds of Eurasian watermilfoil.
Economic damage. The negative impacts on wildlife and fish populations in waterbodies with high densities of Eurasian watermilfoil and the difficulty of motorboating and swimming in infested areas result in recreation-oriented financial losses and the depreciation of shoreline property values (accompanied by a loss of real estate taxes to local economies). It is estimated that milfoil control efforts have cost the United States millions of dollars per year nationwide. Annual control costs in New York state easily exceed half a million dollars per year.
Taxonomy
Eurasian watermilfoil belongs to the watermilfoil family, Haloragaceae, which has two genera in the eastern United States, Myriophyllum L. (10 species, the watermilfoils) and Proserpinaca L. (two species, the so-called mermaid-weeds). All species are submersed herbs inhabiting quiet waters or rooted on muddy shores; all have many finely divided leaves. The species are very similar, resulting in difficulty in identification using only individual specimens or ones without flowers. This has lead to a debate about whether reports of infestations prior to 1940 are actually misidentifications of native species. Myriophyllum spicatum is variable in appearance with long stems, and 12 to 21 leaflet pairs which are limp when out of water. Native M. sibiricum has 5 to 10 leaflet pairs which remain rigid when out of the water. Flowers are arranged on emersed spikes which bear whorls of female flowers basally and whorls of male flowers apically. Female flowers produce four small (2 to 3 mm, 0.08 to 0.16 inch) nutlike fruits .
Biology
Eurasian watermilfoil inhabits ponds and lakes that vary from deep (greater than 100 m, 328 feet) to very shallow (less than a meter, or yard). Waters inhabited may be stagnant, slow-moving fresh, or even slightly brackish. Plants overwinter rooted in the sediment and grow rapidly once favorable warm temperatures are reached in the spring. Because the species tolerates lower water temperatures than most native plants it begins to photosynthesize and grow earlier in the spring than natives, giving the plant the ability to reach the water surface before native plants. The dense canopy over developing native vegetation allows milfoil to out-compete natives for sunlight and space. Flowering typically takes place in early summer and can continue for several months. Although Eurasian watermilfoil produces seed, fragmentation is believed to be the most likely mode of spread in North America. Under unfavorable conditions (high boating traffic, grazing by herbivores or parasites), milfoil may not reach the water surface and won’t flower. M. spicatum has been found to hybridize with native M. sibiricum (producing M. sibricum X spicatum) with an intermediate number of leaf segments. The hybrid plant tends to be more aggressive than the invasive parent species.
Geographic Distribution
Myriophyllum spicatum is native to Europe, Asia, and North Africa. It appears to have been accidentally introduced into North America sometime in the period between the late-1800s and 1940. From the initial points of introduction in the Northeast, M. spicatum has spread to 45 states and at least three Canadian provinces. It has now become a major nuisance species throughout most of the northern US. It is listed as a noxious or otherwise restricted plant in 17 states (in NYS Eurasian watermilfoil is classified as “prohibited”). Milfoil can be spread throughout a waterbody as fragments tangled on boats and trailers or in currents. Furthermore, motor boating and mechanical weed harvesting produce and distribute stem fragments allowing increased propagation. Long distance overland dispersal may be related to the aquarium and aquatic nursery trades.
Figure 3. 2015 U.S. distribution of Eurasian watermilfoil (Map: USDA Natural Resources Conservation Service, plants.usda.gov.
Control
An important aspect of milfoil control is to minimize its spread. Because milfoil is spread overland mostly by human intervention (particularly as a hitchhiker on recreational boats, harvesters, and work barges/boats, it is critical to remove all plant fragments and rinse all equipment that has been in infested waters. The equipment should then be allowed to dry completely before being used in another body of water.
Mechanical Control
Mechanical harvesting of milfoil is used widely used throughout the Northeast and Midwest. Small populations of milfoil around docks, in swimming areas, or near water intakes can be carefully hand-pulled or raked. The best results are found when using multiple harvests per growing season. Care should be taken to prevent breaking off fragments that can float away to start new populations elsewhere. Such localized control can also be undertaken by covering the bed of the waterbody with opaque fabric, thus blocking the light that the plants need to grow.
For use in large areas infested by milfoil, the use of large mechanical harvesters is an option. As in the case of hand-pulling, care should be taken to prevent the movement of fragments to uninfested areas; equipment should be thoroughly cleaned before being moved to other waterbodies. Harvested plant matter can be burned, buried, composted, or by disposed of in landfills.
Eurasian watermilfoil can be killed by dehydration. On managed waterbodies, manipulation of water levels through drawdowns exposing standing biomass and root crown to several weeks of drying time (especially during sub-freezing temperatures).
Mechanical harvester for Eurasian watermilfoil control. Photo: Pelots Bay Association
Chemical Control
A number of chemicals impact the growth and survival of M. spicatum. Amine salts of Endothall (Hyrothol 191®), and Dipotassium Salts of Endothall (Aquathol K®), Diquat dibromide (Reward®), Komeen® have been found to be effective. Some of these herbicides may also affect other non-target rooted submerged plants, including some rushes. Treatment is most effective in still water in the spring while the plant is actively growing.
The amine formulations of 2,4-D granules (Navigate®, Aquakleen®, Aquacide®) are effective on controlling Eurasian watermilfoil and will not damage most non-target grasses. This herbicide method, however, is not appropriate for large unmanageable areas of milfoil.
One lose-dose application (10 µg/ L) of fluridone (brand names Sonar® and Avast!®) applied in the early stages of growth has the potential to provide season-long control of milfoil. However, this application rate causes collateral damage to native vegetation.
Liquid triclopyr (Renovate 3® and Renovate® OTF) can control milfoil without unintended damage to cattails and grasses.
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, particularly the mention of brand names in this profile does not represent a recommendation by NY Sea Grant or Cornell University.
Biological Control
Since the early-1960s, the grass carp, Ctenopharyngodon idella, has been used to reduce the abundance of invasive and nuisance aquatic plants, including Eurasian watermilfoil, in North America. Unfortunately, in many cases grass carp may only eat Eurasian watermilfoil after native plants have been consumed. Effective control of milfoil therefore means the total removal of native aquatic species the fish find more palatable before the grass carp will consume the targeted Eurasian watermilfoil. This may be acceptable if milfoil is the only aquatic plant species in the lake, but due to the substantial negative impacts on native vegetation, grass carp are generally not recommended for control of Eurasian watermilfoil.
For decades, research has evaluated potential insect and pathogen agents for the biological control of Eurasian watermilfoil. Several species of insects have been identified feeding on Eurasian watermilfoil to a damaging degree in North America. Some of these are species to North America while others may have been introduced accidentally from Europe along with introductions of M. spicatum.
The fungus Mycoleptodiscus terrestris has been shown in laboratory research to reduce the biomass of Eurasian watermilfoil significantly and may serve as a possible biocontrol agent.
The North American herbivorous weevil, Euhrychiopsis lecontie, may be associated with recent natural declines in Eurasian milfoil abundance in some lakes in North America. E. lecontei feeds on new growth M. spicatum and may help keep populations under control without concomitant impact on native species. The native midge Cricoptopus myriophylli (Oliver) are also contenders for this recorded damage to milfoil beds.
As many as 20 species of insects appear to feed on M. spicatum on other continents but do not exhibit the host specificity that would be required to make them candidates for milfoil biocontrol.
Among insect species being studied are the North America native weevil Litodactylus leucogaster (Marsham), which attacks the emersed flower spikes of various milfoil species (including non-target species), and the aquatic midge Cricotopus myriophylli (native from New York to British Columbia). While being implicated in the field for contributing to milfoil reduction in BC, the midge does not appear to contribute significantly to declines in lab experiments.
A promising candidate is the naturalized pyralid moth Acentria ephemerella.A. ephemerella is a generalist herbivore which feeds on a variety of aquatic plants. Field evidence indicates it has been associated with declines in Eurasian watermilfoil populations in Ontario, Canada and in New York. In laboratory and in controlled in lake-enclosure experiments, A. ephemerella reduced biomass and plant height and prevented canopy formation.
The North American herbivorous weevil, Euhrychiopsis lecontie. (Photo: Robert L. Johnson, Cornell University, Bugwood.org)The naturalized pyralid moth Acentria ephemerella (Photo: Robert L. Johnson, Cornell University, Bugwood.org)
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
Hydrilla (Hydrilla verticillata), also commonly called water thyme, is a submersed perennial herb. The plant is rooted in the bed of the waterbody and has long stems (up to 25 feet in length) that branch at the surface where growth becomes horizontal and forms dense mats. Small (2 – 4 mm wide, 6 – 20 mm long), pointed, often serrated leaves are arranged around the stem in whorls of 3 to 10. Southern populations are predominantly dioecious female (plants having only female flowers) that overwinter as perennials. Populations north of South Carolina, including populations in New York, are essentially monoecious (having both male and female flowers on the same plant) that set some fertile seed, and depend on tubers for overwintering. These monoecious plants produce female flowers with three translucent petals 10 – 50 mm long by 4 – 8 mm wide, and male flowers with three white to red narrow petals about 2 mm long.
Close-up of Hydrilla. Photo: Chris Evans, River to River CWMA, Bugwood.org
Origin
The dioecious form of Hydrilla is believed to originate from the Indian subcontinent, specifically the island of Sri Lanka, although random DNA analysis also indicates India’s southern mainland as a possible source location. The monoecious form is believed to have arrived on our shores from Korea.
Habitat
Hydrilla can be found infesting freshwater lakes, ponds, rivers, impoundments and canals.
Hydrilla infestation of small lake. Photo: Tim Murphy, University of Georgia, Bugwood.org
Introduction & Spread
The dioecious strain of H. verticillata was imported as an aquarium plant in the early 1950s. Discarded (or intentionally planted ) colonies were found in canals in Miami and Tampa shortly after. The monoecious strain was introduced separately decades later in the Potomac Basin.
Both dioecious and monoecious Hydrilla propagate primarily by stem fragments, although turions (buds) and subterranean tubers also play an important role. The main means of introduction of Hydrilla is as castaway fragments on recreational boats and trailers and in their live wells. New colonies can often be found near boat ramps as such stem pieces become rooted in the substrate (even very, very small fragments can become the start of new populations). Boat traffic through established populations can shatter and spread Hydrilla throughout the waterbody, similar to the spread of Eurasian watermilfoil.
Hydrilla is often a contaminant on popular watergarden plants and may be unwittingly transported and established in private ponds in this manner. As with most invasive aquatic plant species, Hydrilla is a very opportunistic organism and can often be found taking over waters that have had populations of Eurasian watermilfoil chemically removed without a management plan for reestablishing native vegetation.
Impacts
Hydrilla can invade deep, dark waters where most native plants cannot grow. The plant’s aggressive growth (hydrilla’s 20 – 30 foot stems can add up to an inch per day) can spread into shallow water areas and form thick mats that block sunlight to native plants below, effectively displacing the native vegetation of a waterbody. Major colonies of hydrilla can alter the physical and chemical characteristics of lakes:
It is one of the world’s worst aquatic invasive plants
It blocks sunlight and displaces native plants below with its thick, dense surface mats
Stratification of the water column and decreased dissolved oxygen levels can lead to fish kills
The weight and size of sportfish can be reduced when open water and natural vegetation are lost
Waterfowl feeding areas and fish spawning sites are eliminating by dense surface mats
Thick mats of vegation can obstruct boating, swimming and fishing
The value of shorefront property can be significantly reduced, hurting both homeowners and the communities that rely on taxation of shoreline property
In severe infestations, intakes at water treatment, power generation, and industrial facilities can be blocked
Emergent stems and leaves of Hydrilla. Photo: David J. Moorhead, University of Georgia, Bugwood.org
Identification
Hydrilla has pointed, bright green leaves about 5/8 inches long. The leaves grow in whorls of 3 – 10 along the stem, 5 being most common. The margins of the leaves are serrated (toothed). Thin stalks from the stem end in a single, small, floating white flower at the water’s surface. A key identifying feature is the presence of small (up to half inch long), dull-white to yellowish, potato-like tubers which grow 2 to 12 inches below the surface of the sediment at the ends of underground stems. These tubers form at the end of the growing season and serve to store food to allow Hydrilla to overwinter.
Illustration: Cayuga Lake Watershed Network (Rev. October 3, 2012, CCE ISP)Close-up of H. verticillata stem and leaves. Photo: Robert Vidéki, Doronicum Kft., Bugwood.org
Native Lookalikes
Hydrilla is often confused with the common native water weed, Elodea Canadensis, whichhas whorls of 3 smooth-edged leaves as opposed to whorls of 4 to 10 serrated and spined leaves.
Line art: University of Florida Center for Aquatic Plants
Prevention
The best way to help prevent the spread of Hydrilla is to follow basic clean boating techniques:
For All Types of Watercraft:
Be aware of and, if possible, avoid passing through dense beds of aquatic vegetation
Inspect your watercraft, all equipment, and trailers after each use for any plant material
Remove and dispose of all plant matter, dirt, mud and other material in a trash can or above the waterline on dry land well away from where it might get washed back into the lake
Clean and dry all equipment thoroughly before visiting other water bodies (including anything that got wet, such as fishing gear and the family dog)
For Non-Motorized Craft Such as rowing shells, canoes, kayaks, and sailboards:
Open airlocks on shells or air bladders on kayaks after use and allow to dry thoroughly, as plant fragments can survive moist conditions for many days
Around Docks, Launch Sites, and Other Areas:
If plant fragments are piling up around dock areas, use a rake to remove plant material and dispose in the trash
Control
Mechanical harvesting and herbicide spraying are common control methods of controlling Hydrilla. Both are expensive and only moderately effective.
Power weed cutters mow underwater weeds below the water surface and gather them onto a conveyor. The harvesting process is expensive, costing over $1,000 per acre. Because of Hydrilla’s rapid growth, mechanical harvesting needs to be performed several times per growing season. Since the mowing and removal process cannot capture every single fragment of Hydrilla stem and leaf, water and wind currents moving away from the harvest area can easily carry these fragments to uninfested areas of a waterbody and result in new populations taking root.
Chemicals are easier to apply, but also costly. Herbicide spraying works best in small, enclosed bodies of water, and does not work at all in larger bodies the size of a Finger Lake, or in moving water such as a stream, river or canal. Herbicides can also have unintended impacts on native flora, as well. For those reasons, permits for chemical control of Hydrilla are difficult to obtain in New York.
Biological control insects as part of efforts to control Hydrilla have been attempted in Florida with mixed results. Leaf-mining flies from Australia and India and a tuber-feeding weevil from India have been used overseas. The insects released are not native to NY, nor are they currently permitted for release in the State. The use of non-native species to attempt to control another non-native species can be risky if the newly released species out-competes native insects, causing a new invasive species problem. The use of sterile grass carp has been used with some success in small lakes in the southern US but would be impractical in lakes the size of the Finger Lakes.
Another method of dealing with Hydrilla infestations is the control of water levels. Temporary control of Hydrilla has been shown to result from large-scale, long-term water drawdowns. However, since new plants can grow from the buried tubers, regrowth can take place when water levels are allowed to return to normal. Drawdowns also can have negative environmental impacts on native plant species and on fish populations.
Suction harvesting of Hydrilla growth by divers using very strong vacuum hoses can be used to remove Hydrilla from confined areas. However, as with drawdowns, if the underground tubers are not removed by dredging following the suction harvesting, regrowth can take place from the tubers during the next growing season. Further, any fragments that might escape during vacuum activities can float away to root and start new infestations.
The “best”, most effective way to control Hydrilla is the prevention of new Hydrilla infestations.
Eastern US Occurrences
Waterbodies infested with Hydrilla can be found in 70% of Florida’s freshwater drainage basins, making it the most abundant aquatic plant in that state’s waters. Hydrilla is also widespread throughout Alabama; impoundments on the Tennessee River; eastern Mississippi; southeastern Tennessee; southwestern Georgia; South Carolina; eastern North Carolina; in Virginia’s Potomac, Rappahannock, and Appomattox Rivers and into the piedmont, in the tidal freshwater reaches of the Potomac River on the Virginia/Maryland border; along the western and northeastern shores of the Chesapeake Bay, including the Pautuxent River, where it is the most abundant plant species; Pennsylvania (in the Schuylkill River near downtown Philadelphia); eastern Kentucky; in ponds in Delaware; southeastern Connecticut; in a Cape Cod pond in Massachusetts; in southwestern Maine; in New Jersey’s Lower Delaware drainage; Indiana’s Lake Manitou; Wisconsin; and since 2008, in three New York lakes in Suffolk and Orange Counties, and in Cayuga Lake in NY’s Finger Lakes.
Hydrilla can also be found at numerous sites west of the Mississippi River.
Cayuga Lake Inlet Infestation
H. verticillata was detected in the Cayuga Lake Inlet in Ithaca, New York in 2011 by staff of the Cayuga Lake Floating Classroom. A follow-up survey by Robert L. Johnson, a former researcher with the Cornell University Department of Ecology & Evolutionary Biology, now with Racine-Johnson Aquatic Ecologists, located extensive Hydrilla populations in several areas of the Inlet. The Hydrilla appeared to be localized to the Inlet, with no evidence of the plant in Cayuga Lake proper. This was the first detection of Hydrilla in upstate New York. The risk of the plant spreading to the rest of Cayuga Lake and other regional waterbodies in the Finger Lakes region is considered to be substantial. State, regional, and local officials and organizations, along with biologists from Cornell University are developing plans to control, manage, and prevent the spread of the invader, as well as outreach efforts to enlist the public’s help in preventing the plant’s spread.
Profile revised October 3, 2012.
New York Distribution Map
This map shows confirmed observations (green points) submitted to the NYS Invasive Species Database. Absence of data does not necessarily mean absence of the species at that site, but that it has not been reported there. For more information, please visit iMapInvasives.
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