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      Live Snails (Photo by D.L. Gustafson) - Click for full size   Live Snails (Photo by D.L. Gustafson) - Click for full size   Live Snails (Photo by D.L. Gustafson) - Click for full size   Live Snails (Photo by D.L. Gustafson) - Click for full size   Shells (Photo by D.L. Gustafson) - Click for full size   Relaxed snails. Most hydrobiid species must be   Relaxed snails. Most hydrobiid species must be   Field shots show the snails at natural densities (Photo by D.L. Gustafson) - Click for full size   Field shots show the snails at natural densities (Photo by D.L. Gustafson) - Click for full size   Field shots show the snails at natural densities (Photo by D.L. Gustafson) - Click for full size
    Taxonomic name: Potamopyrgus antipodarum (Gray, 1843)
    Synonyms: Hydrobia jenkinsi (Smith, 1889), Potamopyrgus jenkinsi (Smith, 1889)
    Common names: Jenkin's spire shell, New Zealand mudsnail
    Organism type: mollusc
    Potamopyrgus antipodarum is an aquatic snail native to New Zealand that has invaded Australia, Europe, and North America. It can inhabit a wide range of ecosystems, including rivers, reservoirs, lakes, and estuaries. P. antipodarum may established extremely dense populations that can comprise over 95% of the invertebrate biomass in a river, alter primary production, and compete with or displace native mollscs and macroinvertebrates. They can spread rapidly in introduced areas and are able to withstand desiccation, a variety of temperature regimes, and are small enough that many types of water users could be the source of introduction to new areas.
    Description
    Potamopyrgus antipodarum, the New Zealand mudsnail, is a very small, aquatic snail whose elongate shell consists of 5 to 6 dextral, or right handed, whorls. It is often described as horn colored or light to dark brown. It has an operculum that covers its shell aperture. The average length of P. antipodarum is usually 4-6 mm in introduced locations but may reach 12 mm in its native range. Some populations bear a weak keel located mid whorl (Crosier et al, undated; Levri et al, 2007; NZMS Working Group, 2006; Ponder, 1988; Richards et al, 2002; Zaranko et al, 1997).
    Occurs in:
    estuarine habitats, lakes, water courses
    Habitat description
    Potamopyrgus antipodarum is an extremely tolerant species that is capable of inhabiting many aquatic conditions. It colonizes a wide range of habitats including rivers, lakes, streams, estuaries, reservoirs, lagoons, canals, ditches, and even water tanks (Brown et al, 2008; Crosier et al, undated). Reported depths range from 4-25, even 45 meters, but it most often occurs in the littoral zone and moderate depths of around 10 m (Cejka et al, 2008; Zaranko et al, 1997; Grigoorvich et al, 2003). P. antipodarum tolerates a wide range of temperatures, salinities, trophic conditions, water conditions, and current speeds (Gaino et al, 2008; Levri et al, 2007; Crosier et al, undated). It may occupy silt, sand, mud, concrete, vegetation, cobble, and gravel (Crosier et al, undated; Richards et al, 2002). Its densities are reported highest in systems with high primary productivity, constant temperatures, cobble substrate, and constant flow (Richards et al, 2002; Holomuzki & Biggs, 2007), and it thrives in disturbed watersheds (Cejka et al, 2008). Its upper thermal limits are around 28°C and lower limits are around freezing (Crosier et al, undated). It may reproduce at salinities of 0-15 ppt and tolerate 30-35 ppt for short periods of time (Cejka et al, 2008). It can withstand moderate desiccation and drought for several days (National Park Service, undated; Gaino et al, 2008).
    General impacts
    Potamopyrgus antipodarum may establish very dense populations, consume large amounts of primary production, alter ecosystem dynamics, compete with and displace native invertebrates, and negatively influence higher trophic levels. Its ecological plasticity, high competitive ability, high reproductive rate, high capacity for various dispersal methods, and ability to avoid predation make it a formidable colonizer capable of establishing abundant populations with significant effects on ecosystems (Alonso & Castro-Diaz, 2008). P. antipodarum and its impacts are similar to that of the extremely problematic invasive Zebra Mussel (Dreissena polymorpha) (National Park Service, undated).
    P. antipodarum can establish extremely dense populations of tens to hundreds of thousands of individuals per square meter in introduced environments. In Australia densities of 50,000 snails/m2 have been recorded (Ponder 1988; Schrieber et al, 1998). In the United States densities of 200,000, 500,000 and even 800,000 snails/m2 have been recorded in several locations (Davidson et al, 2008; Dorgelo, 1987 in Brown et al, 2008; Crosier et al, undated; Hall et al, 20003; Levri et al, 2007).
    These large populations undoubtedly have significant effects on ecosystems. P. antipodarum can consume up to 75% of gross primary production, dominate secondary production by composing up to 97% of invertebrate biomass, and excreting 65% of total NH4 thereby dominating C and N cycles as in the case of Polecat Creek, Wyoming. Its secondary productivity is one of the highest ever reported (194 g AFDM m-2 yr-1), being 7–40 times higher than that of any macroinvertebrate in Greater Yellowstone area (Hall et al, 2003; Hall et al, 2006; Richards et al, 2002). Such alteration of ecosystems likely results in far reaching cascading ecological impacts (Crosier et al, undated; Davidson et al, 2008; Alonso & Castro-Diaz, 2008). It has also been indicated that it may increase CO2 levels by precipitating calcium bicarbonate to calcium carbonate to produce shells (Chavaud et al, 2003 in NZMS Working Group, 2006).
    P. antipodarum may displace, inhibit growth in, and compete with native invertebrates for resources in introduced locations (Alonso & Castro-Diaz, 2008; Cowie et al, 2009; Davidson et al, 2008; Hall et al, 2006; Kerans et al, 2005). High densities of P. antipodarum were believed to have negative interactions with native macroinvertebrates in several locations in Montana (Kerans et al, 2005). In the Snake River, Idaho, its site of initial introduction in the United States, it is believed to be a major cause of five species of native mollusks recently becoming endangered (Crosier et al, undated). This includes the endangered hydrobiid snail Taylorconcha serpenticola (Richards et al, 2004 in Brown et al, 2008). It is believed to limit absolute growth and the growth rate of the native desert valvata snail (Valvata utahensis) in the Snake River as well (Lysne & Koetsier, 2008). It dominates the Mont Saint-Michael Bay in western France and represented 80% of gastropods collected from all sites (Gerard et al, 2003). Similarly, P. antipodarum made up 83% of the mollusk community in a reservoir near an industrial area in Poland (Lewen & Smolski, 2006). P. antipodarum has been found to significantly inhibit growth in endemic snail Pyrulopsis robusta in Polecat Creek, Wymoing (Riley et al, 2008). A negative correlation has been demonstrated with P. antipodarum and important invertebrate species mayflies, stoneflies, caddisflies, and chironomids (Crosier et al, undated). It has also been to have a negative correlation with native hydrobiid snails in Tasmania (Poner, 1988).
    P. antipodarum directly affects fish by being a poor and mostly un-digestible food source. Although rainbow trout Onchorynchus mykiss and brown trout Salmo trutta were found to feed on P. antipodarum in a study, about 80% of those consumed passed through their system undigested (NZMS Working Group, 2006). Not only does P. antipodarum replace energetic food sources, but it is believed to inflict poor health and reduce survivorship in fish that consume it based the significantly worse condition of fish with P. antipodarum in their guts (Vinsen & Baker, 2008). These direct as well as indirect impacts on fish by P. antiopdarum threaten fisheries in locations where it has established.
    Additionally, P. antiopdarum has fouling potential as it is known to pass through water pipes, emerge from domestic traps, and may block water pipes, meters, or irrigation systems (Ponder, 1988; Cotton, 1942 in Zaranko, 1997; NZMS Working Group, 2006). P. antipodarum has also been found to be infected by blood fluke Sanguinicola sp. in Europe and represents a possible vector to new locations (Gerard & LeLannic, 2003).
    Notes
    Potamopyrgus antipodarum was reported in some locations of Europe as Potamopyrgus jenkinsi by Smith (1989) (Gaino et al, 2008). Non-native populations of P. antipodarum are parthenogenetic and consist almost exclusively of female, clonal individuals. In the United States most western populations are a single clone, with a second in a short section of the Snake River, Idaho, and a third in eastern United States (NZMS Working group, 2006).
    Geographical range
    Native range: New Zealand
    Known introduced range: Austria, Australia, Baltic Sea, Belgium, Black Sea, Canada, Channel Is. (UK), Czech Republic, Denmark, England, Estonia, Finland, France, Germany, Great Lakes (USA), Greece, Italy, Iraq, Japan, Latvia, Lebanon, Lithuania, Netherlands, Northern Ireland, Norway, Poland, Romania, Russian Federation, Scotland, Sea of Azov, Slovakia, Slovenia, Spain, St. Lawrence River, Sweden, Switzerland, Turkey, Ukraine, United States (USA)
    Introduction pathways to new locations
    Agriculture: Commercial movement of aquaculture products, such as live fish or eggs may be an important vector for Potamopyrgus antipodarum spread (Loo et al, 2007a).
    Ignorant possession: The National Park Service (Undated) states that "the rapid spread of P. antipodarum throughout the Madison River watershed may have been assisted by human transport. Mud snails are able to withstand desiccation, a variety of temperature regimes, and are small enough that many types of water users (anglers, swimmers, picnickers, pets) could inadvertently be the mechanism for interbasin transfer of this nuisance species."
    Seafreight (container/bulk): JNCC (2002) states that P. antipodarum "was introduced in drinking water barrels in ships from Australia (Ponder 1988, in JNCC, 2002). The snails were probably liberated while washing or filling water barrels or tanks and, because they can survive in brackish water, they could probably survive liberation into estuarine areas such as the River Thames."
    Ship ballast water: The most frequently cited method of long distance dispersal of Potamopyrgus antipodarum is through ship ballast water (Alonso & Castro-Diaz, 2008).
    Ship/boat hull fouling:
    Stocking: The introduction of Potamopyrgus antipodarum along with fish stocking may be method of its invasion of new locations (Hosea & Finlayson, 2005; Loo et al, 2007b).


    Local dispersal methods
    Boat:
    Consumption/excretion: Potamopyrgus antipodarum is capable of surviving passage through digestion of birds and fish and may be dispersed by them (Cejka et al, 2008).
    Hikers' clothes/boots: The National Park Service (Undated) states that "the rapid spread of P. antipodarum throughout the Madison River watershed may have been assisted by human transport. Mud snails are able to withstand desiccation, a variety of temperature regimes, and are small enough that many types of water users (anglers, swimmers, picnickers, pets) could inadvertently be the mechanism for interbasin transfer of this nuisance species."
    On animals: Potamopyrgus antipodarum may be dispersed to new locations on birds or fish (Alonso & Castro-Diaz, 2008).
    On clothing/footwear: Potamopyrgus antipodarum may b e spread anthropogenically on waders, boots, or angling equipment (Davidson et al, 2008).
    Water currents: Potamopyrgus antipodarum may be dispersed by water currents on floating macrophytes (Alonso & Castro-Diaz, 2008).
    Management information
    Preventative measures: Once Potamopyrgus antipodarum establishes eradication is improbable in most locations and often impractical in those where possible. Prevention of its introduction and containing existing populations is important for minimizing its spread and impacts. Its populations are likely to expand throughout its introduced range. The present distributions of P. antipodarum in North America and Australia specifically are predicted to expand. In North America, it is believed to continue to spread through western watersheds and in the Great Lakes. If it reaches the rivers of the Mississippi basin, it will spread rapidly and abundantly. In Australia it is thought to continue to spread along the east coast and may establish in the southwest if a suitable vector is provided (Loo et al, 2007a).
    Educating anglers, hunters, boaters, aquaculturalists, and the general public about P. anitpodarum, methods of its spread, its potential impacts, and control methods is important. Because its spread to new locations is the result of human activity public awareness about P. antipodarum is necessary. The expansion of present efforts and new initiatives to slow the spread of P. antipodarum by environmental and governmental agencies such as the National Parks Service is essential to conservation (NZMS Working Group, 2006).
    Local and federal governments should also take steps to legally prohibit the importation, possession, and transport of P. antipodarum. In the United States California, Colorado, Kansas, Montana, Utah, Washington, and Wyoming have already done so, while Alaska, Hawaii, Idaho, Nevada, Oregon require prior authorization for its importation, possession, or transport. Colorado and California quarantined and closed fishing access to certain locations in attempts to curb its spread. Alaska, Hawaii, Indiana, Kansas, Montana, Oregon, and Washington have all developed state aquatic nuisance management plans that include P. antipodarum (NZMS Working Group, 2006).
    Transportation via contaminated aquatic equipment, such as wading gear, is a major method of spread of P. antipodarum (Crosier, undated; Davidson et al, 2008; Richards et al, 2004; NZMS Working Group, 2006). Several methods of removing P. antipodarum have been recommended including desiccation, heating, freezing, washing, and chemical treatment. The laying out and drying of equipment at 30ºC for at least 24 hours or at 40ºC for 2 hours has proven effective (Davidson et al, 2008; Richards et al, 2004; Crosier et al, undated). Submerging it in water at about 50 ºC for a few minutes is also effective as P. antipodarum can survive at 43 ºC for short periods (Medhurst, 2003 in NZMS Working Group, 2006). Freezing gear for 6-8 hours will also kill P. anitpodarum (Davidson et al, 2008; Richards et al, 2004; Medhurst, 2003 in NZMS Working Group, 2006). Scrubbing and thoroughly rinsing may effectively remove it as well (Crosier et al, undated). Finally, chemical treatment is also effective. Benzethonium chloride, chlorine bleach, Formula 409, Pine-Sol, ammonia, and copper sulfate all effectively kill P. antipodarum. However, bleach and Pine-Sol were found to damage some materials. The use of copper sulfate, benzethonium chloride, or Formula 409 disinfectant immersion baths or in dry sacks are believed to provide the most acceptable chemical methods of removing P. antipodarum (Hosea & Finlayson, 2005).
    Ballast water and hull fouling is believed to be the most common vector of introducing P. antipodarum to new locations (Alonso & Castro-Diaz, 2008). Adhering to local, federal, and international ballast water regulations such as those provided by GloBallast is essential to reducing the discharge of contaminated ballast water and helping prevent the establishment of P. antipodarum (NZMS Working Group, 2006). Although due to its very small size, it may not be practical to clean P. antipodarum off of large hulls or recreational craft in every instance, promoting information and resources to clean water craft before existing certain contaminated sites would help reduce its spread. Additionally, the cleaning of anchors may also reduce its spread (NZMS Working Group, 2006).

    Physical: Control of P. antipodarum is possible in certain isolated locations such as small lakes, ponds, irrigation canals, and fish hatcheries. Draining waters and allowing substrate to heat and dry completely in the summer or freeze in the winter will kill P. antipodarum. Irrigation canals are routinely shut down for plant control and may be treated for snails as well (NZMS Working Group, 2006). The use of flame throwers on the walls and raceways has been effectively employed in hatcheries (Richards et al, 2004; Dwyer et al, 2003 in NZMS Working Group, 2006). It has also been suggested that barriers such as copper stripping or electrical weirs may limit the movement of P. antipodarum particularly in keeping it from high risk areas (NZMS Working group, 2006).

    Chemical: Chemical treatment of aquatic systems poses risks to surrounding drainages and native species. Small lakes and ponds may be isolated from drainages may isolated from drainages for chemical treatment. Chemical methods used to eradicate P. antipodarum include: Bayer 73 copper sulfate, and 4-nitro-3-trifluoromethylphenol sodium salt (TFM). The only molluscicide known to have been tested against P. antipodarum is Bayluscide (a.i. niclosamide). This test, conducted by Montana Fish, Wildlife, and Parks (FWP), was conducted in small spring creek along the lower Madison River. One hundred percent mortality was achieved after 48 exposure units, which consisted of 1 ppm Bayluscide for 1 hour (Don Skarr, Montana FWP, personal communication in NZMS Working Group, 2006).
    Application of GreenClean® PRO, a non-copper-based algaecide, was found to be an effective means to prevent and possibly eliminate P. antipodarum in the lab. Mortality was 100% within 72 hours of exposure to a 0.5% concentration for 2 and 4 minutes, 1% concentration for 30 seconds, and minimum of 0.33% concentration for 8 minutes. Mortality was also 100%, 48 hours after exposure to a 4% concentration for 2 minutes and 0.55% concentration for 8 minutes. Although effective in the lab, its effectiveness in the remains uncertain (NZMS Working Group, 2006).

    Biological control: Parasites of P. antipodarum are another potential method of control. Studies of the efficacy and specificity of a trematode parasite from its native range as a biological control have demonstrated promising results ((Dybdahl et al. 2005 in NZMS Working Group, 2006; Emblidge and Dybdahl in prep in NZMS Working Group, 2006). Also the parasite Micophallus sp. has been found to highly specific and effective in most genotypes of P. antipodarum including those in the western US (Dybdahl and Lively, 1998 in NZMS Working Group, 2006; Dybdahl & Lively, 1998 in NZMS Working Group, 2006).

    Integrated management: An integrated management and control plan for P. antipodarum should be implemented in locations that are colonized and those that may potentially be invaded. This plan should include preventive measures, public education, monitoring, and appropriate treatment to slow its spread and eradicate where possible and practical. Plans should account for the specific needs of individual locations and follow the guidelines provided by the Aquatic Nuisance Species Task Force (ANSTF) (NZMS Working Group, 2006).

    Nutrition
    Potamopyrgus antipodarum grazes on periphyton, diatoms, and plant and animal detritus (Richards et al, 2002; Alonso & Castro-Diaz, 2008; Brown et al, 2008; Levri et al, 2008).
    Reproduction
    Within its native range Potamopyrgus antipodarum reproduces sexually and asexually while non-native populations are parthenogenetic and consist almost exclusively of triploid females (Alonso & Castro-Diaz, 2008; Lively, undated). Reproduction is ovoviviparous and offspring are brooded by females in a brood pouch until they reach a mobile stage (Alonso & Castro-Diaz, 2008). Broods are reported to range from 20-120 embryos per female and they produce an average of 230 juveniles per year (Richards et al, 2002; Alonso & Castro-Diaz, 2008). P. antipodarum may reproduce year-round in favorable conditions, but the majority of its reproduction occurs in the spring and summer (Crosier et al, undated; Richards et al, 2002).
    Lifecycle stages
    Potamopyrgus antipodarum may live more than a year and has been observed to grow at a rate of up to 0.1 mm/day at 21°C in laboratory conditions (Richards et al, 2002). It may reach sexual maturity in at 3.0-3.5 mm or in about six to nine months (Crosier et al, undated; Richards et al, 2002; Dybdahl & Kane, 2005; Moller et al, 1994 in Alonso & Castro-Diaz, 2008).
    Reviewed by: Dr Sabine Schreiber, Arthur Rylah Institute for Environmental Research Department of Sustainability and Environment. Australia
    Compiled by: National Biological Information Infrastructure (NBII) & IUCN/SSC Invasive Species Specialist Group (ISSG)
    Last Modified: Wednesday, 23 February 2011


ISSG Landcare Research NBII IUCN University of Auckland