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      Zebra mussels attached to aquatic plants (Photo: C. Ramcharan) - Click for full size   Dreissena polymorpha encrusted water meter in Lake Michigan (Photo: M. McCormick, via WikiMedia) - Click for full size   Dreissena polymorpha  (Photo: U.S. Geological Survey Archive, U.S. Geological Survey, Bugwood.org) - Click for full size   Zebra mussels on the lower unit of an inboard/outboard engine (Photo: S. Krynock) - Click for full size   Dreissena polymorpha (Photo: Randy Westbrooks, U.S. Geological Survey, Bugwood.org) - Click for full size
    Taxonomic name: Dreissena polymorpha (Pallas, 1771)
    Synonyms: Mytilus hagenii, Mytilus polymorpha Pallas 1771, Mytilus polymorphus (Pallas), Tichogonia chemnitzii (Rossm.)
    Common names: Dreiecksmuschel (German-Germany), Dreikantmuschel (German-Germany), dreisena (Lithuanian-Lithuania), Eurasian zebra mussel (English), moule zebra (French), racicznica zmienna (Poland), Schafklaumuschel (German-Germany), svitraina gliemene (Latvian-Latvia), tavaline ehk muutlik rändkarp (Estonian-Estonia), vaeltajasimpukka (Finnish-Finland), vandremusling (Danish-Denmark), vandringsmussla (Swedish-Sweden), wandering mussel (English), Wandermuschel (German-Germany, Austria), zebra mussel (English), zebra mussel (Swedish-Sweden), Zebramuschel (German-Germany), Zebra-Muschel (German)
    Organism type: mollusc
    Description
    The shell of D. polymorpha is triangular (height makes 40-60 % of length) or triagonal with a sharply pointed shell hinge end (umbo). The maximum size of D. polymorpha can be 5 centimetres, though individuals rarely exceed 4 cm (Mackie et al. 1989). The prominent dark and light banding pattern on the shell is the most obvious characteristic of D. polymorpha. The outer covering of the shell (the periostracum) is generally well polished, a light tan in colour with a distinct series of broad, dark, transverse colour bands which may be either smooth or zigzag in shape.

    The mussel attaches itself to hard surfaces by byssal threads which are secreted from a byssal gland just posterior to the foot. The byssal threads emerge from the between the valves through a byssal notch along the posterior margin. This byssal hold-fast distinguishes the zebra mussel from all other similar-sized or larger North American freshwater bivalves (McMahon 1990; GSMFC 2005).

    Occurs in:
    estuarine habitats, lakes, urban areas, water courses
    Habitat description
    Zebra mussel larvae are planktonic for 2-4 weeks, prior to beginning their juvenile phase by attaching themselves to substrates by means of byssal threads. Although the juveniles prefer a hard or rocky substrate, they have been known to attach to vegetation (Benson & Raikow 2008). In areas where hard substrates are lacking, such as a mud or sand, zebra mussels cluster on any hard surface available (Benson & Raikow 2008). Given a choice of hard substrates, zebra mussels do not show a preference. Zebra mussels attach to any stable substrate in the water column or benthos including rock, macrophytes, artificial surfaces (cement, steel, rope, etc.), crayfish, unionid clams and each other, forming dense colonies called druses (Benson & Raikow 2008). As adults, they have a difficult time staying attached when water velocities exceed two meters per second (Benson & Raikow 2008). Long-term stability of substrate affects population density and age distributions on those substrates. Within Polish lakes, perennial plants maintained larger populations than did annuals (Stanczykowska & Lewandowski 1993, in Benson & Raikow 2008). Populations on plants also were dominated by mussels less than a year old, as compared with benthic populations; as the mussel colonies grow they sink the macrophytes to which they are attached.

    In their native region zebra mussels will colonise surface standing waters, surface running waters, the littoral zone of inland surface waterbodies, estuaries, brackish coastal lagoons, large estuaries and inland waters, and hard and soft bottom habitats (DAISIE 2006). In their occupied invaded range they will colonise similar habitats with the most typical habitats colonised being lakes, rivers, and estuaries, particularly places where there are firm surfaces suitable for attachment (DAISIE 2006). Zebra mussels tolerate temperatures from -20°C to 40°C; the best growth is observed at 18-20°C (DAISIE 2006). They tolerate brackish waters with salinity up to 7 ppt (DAISIE 2006). They are, however, extremely sensitive to rapid fluctuations in salinity; in the northern Gulf of Mexico, where tidal fluctuations are not great, zebra mussels are found to invade areas with salinities up to 12 ppt, however, they appear unable to tolerate salinities above 12 ppt for any extended period (GSMFC 2005). Zebra mussels prefer moderately productive (mesotrophic) temperate water bodies and occur from the lower shore to depths of 12 m in brackish parts of seas and to 60 m in lakes (DAISIE 2006). They are able to tolerate low oxygen content in water for several days and to survive out of water under cool damp conditions for up to three weeks (DAISIE 2006). Zebra mussel are most abundant in hard waters (30-50 mg Ca L-1) but occur in water with Ca concentrations as low as 12 mg Ca L-1 (Cohen and Weinstein 2001).

    General impacts
    For a detailed account of the environmental impacts of Dreissena polymorpha please read: Dreissena polymorpha Impacts Information. The information in this document is summarised below.

    To date (2002) D. polymorpha has been the most aggressive freshwater invader worldwide (Karayayev et al. 2002). Once introduced, populations of zebra mussel can grow rapidly and the total biomass of a population can exceed 10 times that of all other native benthic invertebrates (Sokolova et al. 1980a; Karatayev et al. 1994a; Sinitsyna & Protasov 1994, in Karayayev et al. 2002)

    Ecosystem Change: Most of the impacts of zebra mussels in freshwater systems are a direct result of their functioning as ecosystem engineers (Karayayev, et al. 2002). An individual zebra mussel can filter one to two liters of water each day; as a result high densities of zebra may cause major shifts in the plankton communities of lakes and rivers. Reductions in phytoplankton numbers and biomass also limit food to fish larvae and other consumers further up the food chain (Birnbaum 2006).

    Modification of Natural Benthic Communities: The introduction of Dreissena is generally associated with increased benthic macroinvertebrate density and taxonomic richness (Ward & Ricciardi 2007). Biodeposition of organic wastes and dense colonization of the benthos by zebra mussels has also substantially altered benthic communities; many invertebrates benefit from the increased food resources and complex habitat, while benthic spawning and foraging fishes may be negatively impacted. Overall gastropod densities increased in the presence of Dreissena, but large-bodied snail taxa tended to decline (Ward & Ricciardi 2007).

    Habitat Alteration: The high consumption of phytoplankton by zebra mussels results in increased water clarity, changing habitat characteristics and ecosystem functions (DAISIE 2006). The dense colonization of soft substrates can impede fish foraging (Beekey et al. 2004), and colonization of hard substrates affects spawning fishes (Marsden & Chotkowski 2001).

    Predation: Zebra mussel populations significantly deplete plankton densities as a result of filter feeding.

    Competition: Suspension-feeding species may experience increased competition for resources in the presence of high zebra mussel densities, as was reflected in the declines of sphaeriid clams in the Hudson River (Strayer, et al. 1998).

    Modification of Nutrient Regime: Zebra mussels may influence ecosystem processes such as nitrogen (N) cycling by increasing denitrification rates (Bruesewitz et al. 2006).

    Threat to Endangered Species: Freshwater mussels (Order Unionoida) are the most imperiled faunal group in North America with 60% of the species considered endangered or threatened (Ricciardi et al. 1998). The zebra mussel represents a new stress to populations of these native mussels as it is a biofouling organism that smothers the shells of other molluscs and competes with suspension feeders for food (Ricciardi, et al. 1998).

    Biofouling: Other mussels serve as substrate for settlement by Dreissena, and are energetically stressed and eventually starve as filter feeding is disrupted (Böhmer et al. 2001, in Birnbaum 2006)

    Economic Impact: Negative economic impacts caused by D. polymorpha include those caused by fouling of intake pipes, ship hulls, navigational constructions and aquaculture cages; the zebra mussel may also reduce angling catches (Gollasch & Leppäkoski 1999; Minchin et al. 2002, in Birnbaum 2006)

    Bioaccumulation: Zebra mussels may bioaccumulate pollutants which may poison animals further up the food chain (DAISIE 2006).

    Uses
    Bioindicator: Due to its sensitivity to anthropogenic influences Dreissena is important as a bioindicator and biomonitoring organism (Franz 1992, in Birnbaum 2006), and quantitative assessments have been conducted regularly since the 1960s in the context of water quality surveys (e.g. in the Rhine) (Schiller 1990, in Birnbaum 2006).

    Products: Crushed shells of the zebra mussel can be used as fertiliser and poultry feed (Birnbaum 2006). Zebra mussels have been used as fishing bait and for fish meal production (DAISIE 2006).

    Notes
    The rapid expansion of the zebra mussel has been linked to its possession of planktonic veliger larvae, byssal threads (for attachment to hard surfaces) and high rates of growth and recruitment (Stanczykowska 1977; Carlton 1993, in Ricciardi Serrouya & Whoriskey 1995b).

    The specific name polymorpha derives from the many variations in shell colour, pattern and shape (Birnbaum 2006).

    Geographical range
    Native range: Native to the drainage basins of the Black, Caspian, Aral and Azov seas (DAISIE 2006; Stanczykowska 1977 in Birnbaum 2006).
    Introduced range: Introduced to north-west Russia, central and western Europe, Scandinavia, Britain, Ireland and North America (DAISIE 2006). During the 19th century the zebra mussel occupied most of inner water systems of western and central Europe, in the 1920s it appeared in Sweden, in the 1960s it was found in alpine lakes around the Alps and reached Italy in 1977, Ireland by 1994 and Spain by 2001 (DAISIE 2006). In 1988 it first appeared in Lake St. Clair and rapidly spread throughout the Great Lakes and large river drainages of North America (DAISIE 2006); it appeared on the west coast in California in 2008 Further range expansions are expected in temperate latitudes of the Northern Hemisphere (DAISIE 2006). Future expansion to South America, South Africa, Australia and New Zealand is possible (DAISIE 2006).
    Introduction pathways to new locations
    Floating vegetation/debris: Zebra mussels attach to floating material and may readily be transported on vegetation or flotsam.
    Pet/aquarium trade: The zebra mussel is possibly introduced into the wild by aquarium dumping.
    Ship ballast water: The main pathways of the expansion in the range of D. polymorpha are through oceanic shipping, in ballast water, and inland navigation, through solid ballast and other cargoes. Inland navigation transport increased since the opening of new waterways between eastern and central Europe at the beginning of the 1800s (Martens 1865, Rebhan 1984, Kinzelbach 1992, Dreyer 1995, Reinhold & Tittizer 1997, Nehring & Leuchs 1999, Gollasch 1996, Orlova 2002, Nehring 2002, in Birnbaum 2006), and within North America (e.g., Marsden & Hauser 2009).
    Ship/boat hull fouling: Zebra mussel adults routinely attach to boat hulls and floating objects and are thus anthropogenically transported to new locations (Benson & Raikow 2008). Humans may spread zebra mussels considerable distances upstream on the hulls of commercial barges (Keevin et al. 1992, in Ricciardi Serrouya & Whoriskey 1995b) and to isolated lakes and rivers through fishing and boating activity (Carlton 1993, McNabb 1993, in Ricciardi Serrouya & Whoriskey 1995b).
    Translocation of machinery/equipment: Results of a study by Ricciardi Serrouya & Whoriskey (1995b) suggest that, given temperate summer conditions, adult Dreissena may survive overland transport (e.g., small trailered boats) to any location within three to five days drive of infested waterbodies.
    Transportation of habitat material: D. polymorpha could be transported with timber or river gravel and overland transport (DAISIE 2006).


    Local dispersal methods
    Aquaculture (local): Larvae may be transported during fish stocking and in bait buckets.
    Boat: The zebra mussel's rapid dispersal throughout the Great Lakes, USA, and major river systems was due to its ability to attach to boats navigating these lakes and rivers (Benson & Raikow 2008).
    Natural dispersal (local): During the pelagic state veligers and post-veligers are transported by currents (DAISIE 2006). Secondary dispersal occurs by the drifting of post-larvae and young adults using byssal and/or mucous threads (Martel 1993, DAISIE 2006).
    On animals: Byssal threads have been an important adapatation for the zebra mussel's success in invading North America (Benson & Raikow 2008). Byssal threads develop in the larvae of some non-dresissenid endemic bivalves and are used to attach to fish gills, there are no endemic freshwater bivalves with byssal adult stages. Speculation exists that waterfowl can disperse zebra mussels, but this has yet to be conclusively demonstrated (Benson & Raikow 2008).
    Other (local): Zebra mussel larvae may be transported on scuba divers' wetsuits, in felt soles of wading boots ,or in scientific sampling equipment.
    Transportation of habitat material (local): Zebra mussel adults attach to aquatic floating plants and may disperse great distances this way (Horvath & Lamberti 1997).
    Water currents: Its rapid dispersal throughout the Great Lakes was also due to the passive drifting of the larval stage (Benson & Raikow 2008).
    Management information
    The following control methods for zebra mussel are potentially useful in certain circumstances (Benson and Raikow 2008):
    • Chemical Molluscicides: Oxidizing (chlorine, chlorine dioxide) and non-oxidizing
    • Manual removal (pigging, high pressure wash)
    • Dewatering/desiccation (freezing, heated air)
    • Thermal (steam injection, hot water 32oC)
    • Acoustical vibration
    • Electrical current
    • Filters/screens
    • Coatings: toxic (copper, zinc) and non-toxic (silicone-based)
    • Toxic constructed piping (copper, brass, galvanized metals)
    • CO2 injection
    • Ultraviolet light
    • Anoxia/hypoxia
    • Flushing
    • Biological (predators, parasites, diseases)

    Preventative measures: Preventing overseas transfer can only be achieved by mid-ocean exchange or by suitable disinfection of ballast water (DAISIE 2006). Certain guidelines and regulatory instruments may be applied in areas where the species does not yet occur (Gollasch 2006). For further details see the Ballast Water Management Convention of the International Maritime Organization (www.imo.org) and the Code of Practice for the Introduction and Transfer of Marine organisms of the International Council for the Exploration of the Sea (www.ices.dk).
    Appropriate control measures (inspection, removal of attached mussels, drying, etc.) should be taken to minimise risk of inoculation by transfer of boats, fishing gears, etc (DAISIE 2006). Applying copper based anti-foulant coatings in new facilities may offer protection from Dreissena polymorpha. The use of retrofitted screens can be effective but such screens are difficult to apply to existing pipelines (Aldridge et al. 2006).

    Physical: Physical removal using high-pressure water jets is feasible on easily accessed industrial facilities (Aldridge et al. 2006). Larvae suffer total mortality after exposure to ultrasonic vibration (22 to 800 kHz) for 3 minutes (Schalekamp 1971, in Birnbaum 2006), but the technical effort involved is prohibitive.

    Chemical: Many chemicals will kill zebra mussels but the suitability of a particular chemical is determined by considerations of effect on water quality, residual concentrations, byproducts, cost and practicality. Chemicals which have proven moderately successful include molluscicides (such as Bayer 73; Birnbaum 2006), chloramines, chlorine dioxide, ozone, hydrogen peroxide, potassium permanganate, pH adjustment, and inorganic salts. Chlorination remains the only widespread method used. It must be dosed continuously for up to 3 weeks to achieve complete elimination, though dosing for 2-3 days is sufficient to remove the majority of attached mussels.
    Microencapsulation of toxins in particles that are edible to zebra mussels has the potential to overcome the rejection and valve-closing response generally seen when zebra mussels are exposed to toxic substances. The active ingredient used is potassium chloride, which is not lethal to most organisms, including fish, at low doses but which is particularly toxic to freshwater bivalves (Aldridge et al. 2006). Another emerging control for D. polymorpha is the use of endocannabinoids, anandamide and other compounds which have been tested to inhibit zebra mussel byssal attachment. These naturally occurring and synthetic cannabinoids can serve as non-toxic efficacious zebra mussel anti-foulants (Angarano et al. 2009).

    Biological control: Large-bodied molluscivores such as common carp, freshwater drum, and channel catfish can limit zebra mussel numbers in coastal wetlands. Densities of other molluscs were not affected, suggesting that fish can have a greater impact on numbers of attached zebra mussels than other benthic molluscs (Bowers & DeSzalay, 2007). Known predators also include roach, eel, sturgeon, diving ducks, crayfish and muskrats (Molloy et al., 1997).

    Nutrition
    Zebra mussels filter a wide range of size particles, but select only algae and zooplankton between 15 and 400 microns. Larval stages of the mussel feed on bacteria.
    Reproduction
    Zebra mussels have separate sexes, usually with a 1:1 ratio; fertilisation takes place externally (DAISIE 2006). Synchronised spawning occurs once mussels are greater than 8 mm (or females in their second year) and is influenced by water temperatures (DAISIE 2006). A mature female may produce one million eggs per year (DAISIE 2006). Spawning begins at 12 to 15ºC and is optimal at 14 to 16ºC or 18 to 20ºC (depending on sources) and may take place over a period of three to five months (DAISIE 2006; Benson & Raikow 2008). In natural ecosystems oogenesis occurs in autumn, with eggs developing until release and fertilization in spring; in areas of warm water or where the thermal regime has been altered, reproduction can occur continually throughout the year (Benson & Raikow 2008). Eggs are expelled by the females and fertilized outside the body by the males; over 40 000 eggs can be spawned in a reproductive cycle and up to one million in a spawning season (Benson & Raikow 2008).
    Lifecycle stages
    Fertilised eggs hatch into trocophores (40-60 microns, 1 to 2 days), which develop within a day into a free-swimming planktonic veliger. Veligers develop from a d-shaped to umbonal morphology, and remain planktonic for up to 4 weeks. Optimal temperature for larval development is 20 to 22oC (Benson & Raikow 2008). Larvae normally disperse by being passively carried downstream with water flow (Benson & Raikow 2008). The larvae develop into their juvenile stage once they have reached about 350 microns in size by settling to the bottom where they crawl about by means of a foot, searching for suitable substratum (Benson & Raikow 2008). They then attach themselves to substrates by means of a byssus, a cluster of threads produced by an external organ near their foot (Benson & Raikow 2008). They may mature within the first year of life under optimal conditions; maturity in the second year is more usual. Once attached, the life span of D. polymorpha is variable, but can range from 3 to 9 years (Benson & Raikow 2008). Adult mussels can voluntarily detach and move around the substrate to seek alternate locations.
    This species has been nominated as among 100 of the "World's Worst" invaders
    Reviewed by: J. Ellen Marsden, Rubenstein School of Environment and Natural Resources, University of Vermont,Burlington, USA.
    Compiled by: Profile revision: National Biological Information Infrastructure (NBII) & IUCN/SSC Invasive Species Specialist Group (ISSG)
    Last Modified: Tuesday, 22 September 2009


ISSG Landcare Research NBII IUCN University of Auckland