Limax maximus (leopard slug)
- Summary of Invasiveness
- Taxonomic Tree
- Notes on Taxonomy and Nomenclature
- Distribution Table
- History of Introduction and Spread
- Risk of Introduction
- Habitat List
- Hosts/Species Affected
- Growth Stages
- List of Symptoms/Signs
- Biology and Ecology
- Natural enemies
- Notes on Natural Enemies
- Means of Movement and Dispersal
- Plant Trade
- Wood Packaging
- Economic Impact
- Environmental Impact
- Threatened Species
- Social Impact
- Risk and Impact Factors
- Detection and Inspection
- Similarities to Other Species/Conditions
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- Limax maximus Linnaeus
Preferred Common Name
- leopard slug
Other Scientific Names
- Heynemannia maximus (L.)
- Krynickillus mamelianus Bourguignat
- Limacella parma Brard
- Limax abrostolus Bourguignat
- Limax antiquorum Férussac
- Limax bocagei da Silva e Castro
- Limax carbonarius albanicus Jaeckel
- Limax cellarius Dezallier d’Argenville
- Limax cinereus Müller
- Limax maculatus Nunneley
- Limax maximum
- Limax maximus altenai Grossu & Lupu
- Limax pardalis R.T. Lowe
- Limax sylvaticus Draparnaud
- Limax vulgaris Moquin-Tandon
- Milax maximus (L.)
International Common Names
- English: European giant garden slug; great gray garden slug; great grey slug; great slug; large garden slug; large grey slug; spotted garden slug; spotted leopard slug; tiger slug
- Spanish: babosa atigrada; gigante limacido grisaco; gran babosa gris; gran limacido grisaceo
- French: grande limace cendree; grande limace grise; grande limace jaune; grande limace tachetee; limace cendree; limace léopard
Local Common Names
- Albania: ligaveci i madh; ligaveci tiger
- Chile: babosa grande del jardín
- Croatia: veliki balavac
- Denmark: gra pantersnegl; grasnegl; pantersnegl, skov-; skovpantersnegl; skov-pantersnegl; stor kjølsnegl
- Faroe Islands: leopardsnigil
- Germany: Grosse Kellerschnecke; Grosse Nacktschnecke; Grosse Wegschnecke; Grosser Schnegel; Pantherschnecke; Schnecke, Grosse Keller-; Tigerschnegel
- Iceland: ardussnigill
- Italy: limaccia massima; lumacone maggiore
- Netherlands: aardslak; grote aardslak
- Norway: boakjølsnegl
- Poland: pomrów wielki
- Sweden: stor trädgardsnigel; traedgardsnigel, stor
- Switzerland: Pantersnigel
- USA: great gray garden slug
- LIMXMA (Limax maximus)
Summary of InvasivenessTop of page
L. maximus is a large slug, not widely recognized as invasive, but often viewed as a plant pest. It has become widely distributed through human agencies, continues to spread in at least some regions, and has often been regarded as a pest of cultivated plants and mushrooms. Herbivory has been recorded on some native plant species, and it can contribute significantly to leaf litter decomposition. It is an intermediate host of nematodes parasitic in mammals, and known to interact with some native mollusc species. L. maximus has not, however, been recognized as threatening ecosystems, habitats or species or having major economic consequences.
Most concern about L. maximus has been expressed in the Hawaiian Islands, where the species has established in upland forests and recent experimental work suggests significant effects on seedling recruitment for forest trees (Joe, 2006; Joe and Daehler, 2008) and on leaf litter decomposition rates (Meyer et al., 2013). The implications are that these effects may lead to ecosystem-level changes, but definitive evidence for this is presently lacking.
There is also developing interest in possible ecosystem consequences of L. maximus [and other adventive slugs] in forests of North America. Recent experimental work suggests effects on seedling recruitment for forest trees, especially balsam fir (Abies balsamea) but definitive evidence for ecosystem-level changes is presently lacking.
L. maximus is not listed as threatened in any part of its native range, reflecting the continuing abundance of the species in both natural and modified habitats.
Taxonomic TreeTop of page
- Domain: Eukaryota
- Kingdom: Metazoa
- Phylum: Mollusca
- Class: Gastropoda
- Subclass: Pulmonata
- Order: Stylommatophora
- Suborder: Sigmurethra
- Unknown: Limacoidea
- Family: Limacidae
- Genus: Limax
- Species: Limax maximus
Notes on Taxonomy and NomenclatureTop of page
The genus Limax comprises over 30 species of large, terrestrial slugs (type species Limax maximusLinnaeus, 1758, by subsequent designation of Férussac, 1819). The genus is native to the European continent (Wiktor and Likharev, 1979; Wiktor, 1996, 2001), with hotspots of diversity in the Mediterranean area (Lessona and Pollonera, 1882; Wiktor, 2001), European Alps (e.g. Simroth, 1885, 1901, 1910; Heynemann, 1905; Hesse, 1926; Simroth and Hoffmann, 1928; Alzona, 1971; Boato et al., 1989), and the Balkans (Rähle, 1976; Wiktor, 1983, 1996). Many of the species are poorly known (Nitz et al., 2009) and that there is often considerable intraspecific variation in body pigmentation has led to the introduction of numerous species-group names and subsequent extensive synonymies.
Only one species in the genus, L. maximus, has been widely distributed by human activities and is considered invasive in at least some areas. The species formerly known as Limax flavus Linnaeus has also been distributed to many parts of the world by human activities, but is now placed in the genus Limacus (see Barker, 1999).
The species now known as Limax maximus was described by Lister (1678), Linnaeus (1745, 1746), Dézallier d'Argenville (1757) and others in published works rejected for the purposes of zoological nomenclature (i.e. pre-Linnean, 1758). Limax maximus was formally described by Linnaeus (1758) by reference to Lister (1678; who gave a detailed description of Limaxcinereus maximus maculatus and striatus from England), and two of his own publications mentioning Limax cinereus maculatus from Sweden (Linnaeus, 1745, 1746). The type locality of L. maximus has long been considered as Sweden but this had not been firmly established. With the designation of a lectotype by Proschwitz and Falkner (2007), based on the published illustrations by Lister (1685: pl., fig. 2), the type locality was established as near York, Yorkshire, England.
DescriptionTop of page
Slug – animal without an external shell. Large, with a saddle-like mantle shield that overs only the anterior part of the body; containing a vestigial shell as an oval plate. Mantle covered with black spots and mottles, back with black mottles and broken bands. Pneumostome or breathing pore (the opening to the lung) – in right posterior margin of mantle shield.
Species described in detail by Quick (1949, 1960), Wiktor (1973, 1983, 1996), Likharev and Wiktor (1980), Barker (1999), and many other authors. See Barker (1999) for terminologies.
Body morphology: Slugs up to 180 mm, rarely to 200 mm long; specimens 100 mm long usually mature. Mantle about 0.3 of body length. Hind body distinctly keeled. Body yellowish white or grey to brown; back usually with 6, 4, or 2 dark (tan, black to navy blue-black) bands, these frequently interrupted to form a more or less coarsely spotted pattern; mantle irregularly spotted or marbled with dark pigment (tan, black to navy blue-black). Tentacles uniformly vinous brown. Sole uniformly pale. Mucus clear. Locomotion by muscular pedal waves generated from posterior of sole. Genital orifice immediately posterior to right ocular peduncle.
Shell: Thin, shiny white, oblong-oval, weakly convex plate, up to 14 X 6.7 mm; dorsal surface with distinct growth lines. Protoconch vestigial, non-protruding, situated asymmetrically at left side near posterior margin. An organic sheet surrounding the fresh shell. Shell often persists for several months or years in the environment after death of the animal.
Reproductive system: Hermaphrodite. Ovotestis elongate, reaching apex of body cavity or almost so, embedded in lobes of the digestive gland. Hermaphrodite duct long, at first straight and slender, then wider and convoluted before narrowing to talon, which is deeply embedded in albumen gland. Female part of spermoviduct folded and voluminous. Free oviduct long and mostly slender, but dilated at entry into atrium. Bursa copulatrix reservoir small, oval, on a short duct opening to base of phallus, very close to atrium. Vagina absent. Prostatic gland fused to female oviduct proximally, free anteriorly. Vas deferens thin, opening at apex of phallus adjacent to insertion of retractor muscle. Phallus cylindrical, its length half or more that of body, strongly convoluted; lacking a caecum; internally with a fold expanded proximally into a comb. Atrium short. Phallus retractor short, arising from left margin of diaphragm in posterior part of pallial complex.
Digestive tract: Jaw about 3.5 mm wide, with a prominent medial projection and transversely scored with fine striae.
Radular ribbon comprising about 150 rows of teeth, each near the formula 20+50+C+50+20. Central tooth tricuspid, with a prominent mesocone flanked on either side by small, weak ectocones. First lateral tooth usually tricuspid, with prominent mesocone flanked by small endocone and ectocone. Lateral teeth with mesocone progessively more slender and elongate towards outer lateral field of radula; ectocone generally absent in 2nd or 3rd lateral tooth; endocone persisting to perhaps the 20th tooth. Marginal teeth markedly smaller than lateral teeth, with mesocone outer edge somewhat serrated.
Buccal mass spheroidal. Oesophagus quickly expanding to large crop, extending to about 0.6 of body cavity. Stomach a simple curvature, with 2 ducts to digestive gland. Intestine arising from left lateral aspect of stomach, running directly forwards to pass over anterior aorta, then producing a short posterior loop overlying crop before again running forwards to pass over stem of cephalic retractor, and finally producing a long, posteriorly directed loop along right side of body cavity before running to anus.
Pallial complex: Located in posterior part of mantle. Kidney bean-shaped, its longest axis somewhat obliquely positioned relative to body axis, partially enclosing heart on its left side. Secondary ureter separating from left posterior part of kidney and describing an arc to right anterior quarter of pallial complex, where it inflates to form an elongate, tubular urinary bladder. Heart in left anterior quarter of pallial complex, with ventricle directed posteriorly and to the right; aortic stem short. Lung with a well-developed vascular network.
Free muscle system: Cephalic retractor attached on dorsal body wall a little posterior to pallial complex and passing forwards before dividing into left and right tentacle retractors; buccal retractor arising as a branch from stem or from left tentacle retractor; second anterior loop of intestine passing over cephalic retractor stem near its origin.
Central nervous system: Located in anterior of body cavity, immediately behind buccal mass. Cerebral ganglia united by a short but distinct commissure. Cerebropedal connectives short, their length less than width of cerebral ganglia. Pleural ganglia markedly closer to pedal ganglia than to cerebral ganglia. Visceral chain compact but with pleural, parietal, and visceral ganglia distinct; visceral ganglion to right of median plane.
Essentially miniatures of the adults in body form, but lacking fully developed reproductive organs. The hatchlings are pale, translucent grey, with a faint band visible on the mid dorsum and vinous ocular peduncles and inferior tentacles. Within 3 weeks the other body bands have appeared and begin breaking up, and the mantle becomes increasingly spotted or mottled.
Varying greatly in size about a mean of 5.0 × 5.5 mm, are soft, translucent, and amber-coloured. They are laid in clusters of 20-100, and hatch in about 1 month in the field (approx. 14 days at 18-20°C: Prior, 1983).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|China||Present||Present based on regional distribution.|
|-Guangdong||Present||Introduced||Yang et al., 2012|
|India||Present||Present based on regional distribution.|
|-Himachal Pradesh||Present||Introduced||Godwin-Austen, 1908|
|Israel||Absent, formerly present||Introduced||Dautzenberg, 1894||Listed by Dautzenberg (1894) as collected at El Bireh, but species not listed by Miens (1982) Roll et al. (2009) or Heller (2009)|
|Japan||Present||Introduced||Not invasive||Mito and Uesugi, 2004|
|-Honshu||Localised||Introduced||Not invasive||Azuma, 1982; Hasegawa et al., 2009|
|Turkey||Widespread||Native||Not invasive||Rähle, 1992b; Schütt, 2005|
|Algeria||Widespread||Native||Not invasive||Taylor, 1902-1907; Forbes, 1838; Quick, 1960|
|Egypt||Present||Native||Not invasive||Abdel-Haleem, 2014|
|Morocco||Present||Native||Not invasive||Dakkak and Cabaret, 1984; Dakkak and Ouhelli, 1988|
|Saint Helena||Present||Introduced||Not invasive||GBIF, 2015|
|South Africa||Localised||Introduced||1898||Not invasive||Melvill and Ponsonby, 1898; Herbert and Kilburn, 2004|
|-Canary Islands||Present||Introduced||Not invasive||Quick, 1960|
|Canada||Present||Present based on regional distribution.|
|-British Columbia||Widespread||Introduced||1947||Not invasive||Quick, 1960; NatureServe, 2014||Pest in Fraser Valley in 1947 (Glendenning, 1947) and Vernon district in 1960 (Arnott & Arrand, 1961)|
|-Newfoundland and Labrador||Present||Introduced||1940||Not invasive||Brooks and Brooks, 1940; NatureServe, 2014|
|-Nova Scotia||Present||Introduced||Not invasive||Davis, 1992; NatureServe, 2014|
|-Ontario||Present||Introduced||1904||Not invasive||Latchford, 1904; NatureServe, 2014|
|-Quebec||Present||Introduced||Not invasive||Pilsbry, 1948|
|Mexico||Present||Introduced||1926||Not invasive||Baker, 1930|
|USA||Present||Introduced||1857||Not invasive||Quick, 1952|
|-Alabama||Present||Introduced||Not invasive||Hubricht, 1965; Texas Invasive Species Institute, 2004|
|-Alaska||Present||Introduced||Not invasive||Ferguson and Knight, 2010|
|-Arizona||Present||Introduced||Not invasive||Bequaert and Miller, 1973; Dundee, 1974|
|-California||Widespread||Introduced||1890||Not invasive||Orcutt, 1890; McDonnell et al., 2009; NatureServe, 2014||San Diego (Orcutt, 1890); Oakland by 1896, Los Angeles by 1901, Monterey by 1902, Pasadena by 1896 (Vanatta, 1904; Pilsbry, 1948)|
|-Colorado||Widespread||Introduced||1918||Not invasive||Henderson, 1918; NatureServe, 2014||Boulder, by 1918 (Henderson, 1918). Uncommon, but present in natural habitat|
|-Connecticut||Present||Introduced||Not invasive||Chichester and Getz, 1969; Texas Invasive Species Institute, 2004|
|-Delaware||Present||Introduced||Texas Invasive Species Institute, 2004; Project Noah, 2013|
|-District of Columbia||Present||Introduced||1937||Not invasive||GBIF, 2015|
|-Florida||Present||Introduced||Not invasive||GBIF, 2015|
|-Georgia||Present||Introduced||Not invasive||Texas Invasive Species Institute, 2004|
|-Hawaii||Widespread||Introduced||1920||Invasive||Williams, 1931; Joe and Daehler, 2008; NatureServe, 2014||Oahu; Maui; Hawaii; Waiahinu; Published records indicates first presence in Hawaii in 1931. However, earlier collected specimens are represented in museum collections|
|-Illinois||Present||Introduced||1901||Not invasive||Baker, 1901; Dundee, 1974; Texas Invasive Species Institute, 2004|
|-Indiana||No information available||Introduced||1926||Not invasive||Thompson, 1927; Texas Invasive Species Institute, 2004; Project Noah, 2013|
|-Iowa||Present||Introduced||Texas Invasive Species Institute, 2004|
|-Kansas||Present||Introduced||Not invasive||Leonard, 1959; Texas Invasive Species Institute, 2004|
|-Kentucky||Present||Introduced||Not invasive||Branson and Batch, 1969; NatureServe, 2014|
|-Louisiana||Present||Introduced||Not invasive||GBIF, 2015||New Orleans|
|-Maine||Widespread||Introduced||1909||Not invasive||Lermond, 1909; NatureServe, 2014||Bar Harbor (Lermond, 1909)|
|-Maryland||Widespread||Introduced||1894||Not invasive||Pilsbry, 1948; Dundee, 1974; NatureServe, 2014|
|-Massachusetts||Widespread||Introduced||1882||Not invasive||Frandsen, 1901; Pilsbry, 1948; Dundee, 1974|
|-Michigan||Widespread||Introduced||Not invasive||Pilsbry, 1948; NatureServe, 2014|
|-Minnesota||Present||Introduced||Texas Invasive Species Institute, 2004|
|-Mississippi||Present||Introduced||Not invasive||Hubricht, 1977||Hinds and Lauderdale counties|
|-Missouri||Present||Introduced||Not invasive||Hubricht, 1972; Dundee, 1974; Texas Invasive Species Institute, 2004|
|-Montana||Present||Introduced||Not invasive||Texas Invasive Species Institute, 2004; Hendricks, 2009; NatureServe, 2014|
|-Nebraska||Present||Introduced||Not invasive||Barger and Hnida, 2008|
|-New Hampshire||Present||Introduced||Texas Invasive Species Institute, 2004|
|-New Jersey||Widespread||Introduced||1889||Cockerell, 1889; Dirrigl and Bogan, 1996; NatureServe, 2014|
|-New York||Widespread||Introduced||1870||Not invasive||Pilsbry, 1948; Wurzinger, 1975; NatureServe, 2014||Long Island by 1870 (Pilsbry, 1948)|
|-North Carolina||Widespread||Introduced||1951||Not invasive||Hubricht, 1970; Dourson and Dourson, 2006; NatureServe, 2014||First published record evidently Hubricht (1970). However, earlier collected specimens are represented in museum collections (e.g. ANSP A1019, collected 1930)|
|-Ohio||Widespread||Introduced||1882||Not invasive||Pilsbry, 1948; Dundee, 1974; Texas Invasive Species Institute, 2004|
|-Oklahoma||Widespread||Introduced||Not invasive||Branson, 1962; Dundee, 1974; Texas Invasive Species Institute, 2004|
|-Oregon||Widespread||Introduced||1898||Not invasive||Henderson, 1929; NatureServe, 2014||Published records indicate first presence by 1929. However, earlier collected specimens are represented in museum collections (e.g. ANSP 72498 A3124c, collected JG Malone 1898)|
|-Pennsylvania||Widespread||Introduced||1867||Not invasive||Tryon, 1867; Dundee, 1974; NatureServe, 2014||Philadelphia by 1867|
|-Rhode Island||Widespread||Introduced||1868||Not invasive||Gould and Binney, 1870; Pilsbry, 1948; NatureServe, 2014||Newport by 1868|
|-South Carolina||Present||Introduced||Texas Invasive Species Institute, 2004|
|-South Dakota||Present||Introduced||Not invasive||Frest and Johannes, 2002; NatureServe, 2014|
|-Tennessee||Widespread||Introduced||Not invasive||Lutz, 1950; Dourson and Dourson, 2006; NatureServe, 2014|
|-Texas||Present||Introduced||1886||Not invasive||Pilsbry, 1948; Texas Invasive Species Institute, 2004|
|-Utah||Present||Introduced||1929||Not invasive||Pilsbry, 1948; NatureServe, 2014|
|-Vermont||Present||Introduced||Texas Invasive Species Institute, 2004|
|-Virginia||Present||Introduced||1889||Not invasive||Cockerell, 1889; Dundee, 1974; NatureServe, 2014|
|-Washington||Present||Introduced||1930||Not invasive||Dundee, 1974; NatureServe, 2014||First published record evidently Dundee (1974). However, earlier collected specimens are represented in museum collections (e.g. ANSP 162862 A3157, collected AJ Hanson 1930)|
|-West Virginia||Present||Introduced||Texas Invasive Species Institute, 2004|
|-Wisconsin||Localised||Introduced||1911||Not invasive||Texas Invasive Species Institute, 2004; Jass, 2007; NatureServe, 2014||First published record Milwaukee 1911 (Jass 2007)|
Central America and Caribbean
|El Salvador||Unconfirmed record||Introduced||Schwartz et al., 1978; Huezo and Lainez, 1986||Reputedly from higher elevation areas but may be based on misidentifications|
|Honduras||Unconfirmed record||Introduced||Schwartz et al., 1978; Schoonhoven et al., 1982||Reputedly from higher elevation areas but may be based on misidentifications|
|Argentina||Localised||Introduced||Not invasive||Hylton Scott, 1963; Virgillito and Miquel, 2013||Neuquén (Hylton-Scott 1963)|
|Brazil||Present||Introduced||Not invasive||Boffi, 1979; Simone, 2006|
|-Parana||Present||Introduced||Not invasive||Graeff-Teixera et al., 1993|
|-Rio Grande do Sul||Present||Introduced||Not invasive||Agudo-Padrón, 2009|
|-Santa Catarina||Present||Introduced||Not invasive||Agudo-Padrón, 2011|
|Chile||Widespread||Introduced||Not invasive||Stuardo and Vega, 1985; Landler and Nuñez, 2012|
|Colombia||Present||Introduced||Not invasive||Schoonhoven and Cardona, 1980||Tenjo (2,587 m (8,488 ft))|
|Albania||Widespread||Native||Not invasive||Bank, 2011a; Jaeckel, 1954|
|Andorra||Present||Native||Not invasive||Bank, 2011c|
|Austria||Widespread||Introduced||Not invasive||Bank et al., 2002a; Taylor, 1902-1907|
|Belarus||Widespread||Introduced||Not invasive||Likharev and Rammel'meier, 1952; Zemoglyadchuk, 2009|
|Belgium||Widespread||Native||Not invasive||Bank et al., 2002b; Taylor, 1902-1907|
|Bosnia-Hercegovina||Present||Native||Not invasive||Bank, 2011a; Wiktor and Jurkowska, 2007|
|Bulgaria||Widespread||Native||Not invasive||Jurinic, 1906; Irikov and Eröss, 2008|
|Czech Republic||Widespread||Introduced||Not invasive||Bank et al., 2002c; Juricková et al., 2001; Dvorák, 2005|
|Denmark||Present||Introduced||1868||Not invasive||Bank et al., 2002d; Malm, 1868; Ferdushy et al., 2009|
|Estonia||Present||Introduced||Bank et al., 2002e|
|Faroe Islands||Present||Introduced||2003||Jensen et al., 2014|
|Finland||Present||Introduced||1886||Esmark, 1886; Routio and Valta, 2009|
|France||Widespread||Native||Moquin-Tandon, 1855; Falkner et al., 2002|
|-Corsica||Present||Not invasive||Simroth, 1900; Holyoak, 1983|
|Germany||Widespread||Introduced||Not invasive||Bank, 2011b; Taylor, 1902-1907; Ludwig et al., 2015||Including Helgoland Island (Caspers, 1942)|
|Greece||Widespread||Native||Not invasive||Taylor, 1902-1907; Wiktor and Jurkowska, 2007|
|Hungary||Widespread||Introduced||1892||Not invasive||Taylor, 1902-1907; Zoltán and András, 2001|
|Iceland||Present||Introduced||1997||Armitage and McMillan, 1964; Ólafsson, 2012|
|Ireland||Widespread||Native||Not invasive||Bank et al., 2002g; Scharff, 1891|
|Italy||Widespread||Native||Not invasive||Betta Ede, 1853; Alzona, 1971|
|Latvia||Widespread||Introduced||Not invasive||Bank et al., 2002i; Stalažs et al., 2014|
|Liechtenstein||Widespread||Native||Not invasive||Bank, 2011e; Trüb, 1988|
|Lithuania||Widespread||Introduced||Not invasive||Bank, 2011f; Šatkauskiene, 2001|
|Luxembourg||Present||Native||Not invasive||Bank et al., 2002k|
|Moldova||Present||Introduced||Likharev and Rammel'meier, 1952; Balashov et al., 2013|
|Monaco||Present||Native||Not invasive||Caziot, 1910|
|Montenegro||Present||Wiktor and Jurkowska, 2007|
|Netherlands||Widespread||Native||Not invasive||Bank et al., 2002i; Taylor, 1902-1907|
|Norway||Present||Introduced||1870||Not invasive||Esmark, 1886; Gederaas et al., 2012|
|Poland||Widespread||Introduced||Slosarski, 1877; Wiktor and Jurkowska, 2007|
|Portugal||Widespread||Native||Not invasive||Bank, 2011c; Morelet, 1845; Rodriquez and Hermida, 1993|
|-Azores||Widespread||Introduced||1860||Not invasive||Morelet, 1860; Cunha et al., 2005|
|-Madeira||Widespread||Introduced||1878||Wollaston, 1878; Seddon, 2008|
|Romania||Widespread||Native||Not invasive||Grossu and Lupu, 1960; Wiktor and Jurkowska, 2007|
|Russian Federation||Present||Present based on regional distribution.|
|-Central Russia||Present||Introduced||Not invasive||Simroth, 1898; Sysoev and Schileyko, 2009|
|-Southern Russia||Present||Not invasive||Taylor, 1902-1907; Sysoev and Schileyko, 2009|
|Serbia||Present||Native||Not invasive||Wiktor and Jurkowska, 2007|
|Slovakia||Widespread||Introduced||Not invasive||Šteffek, 1978; Cejka et al., 2007|
|Slovenia||Present||Introduced||Not invasive||Bole and Slapnik, 1997|
|Spain||Widespread||Native||Not invasive||Bank, 2011c; Graells, 1846; Wiktor and Jurkowska, 2007|
|-Balearic Islands||Present||Not invasive||Bank, 2011c; Castillejo and Rodríguez, 1991|
|Sweden||Present||Introduced||Not invasive||Bank et al., 2002p; Proschwitz Tvon, 1988|
|Switzerland||Present||Introduced||Not invasive||Taylor, 1902-1907|
|UK||Widespread||Native||Not invasive||Taylor, 1902-1907; Meiklejohn, 1973; Anderson, 2005||As far north as Caithness in Scotland (Meiklejohn, 1973)|
|-Channel Islands||Widespread||Native||Not invasive||Duprey, 1876; Chatfield, 1975|
|Ukraine||Present||Introduced||Not invasive||Likharev and Rammel'meier, 1952; Sysoev and Schileyko, 2009|
|-New South Wales||Widespread||Introduced||Not invasive||Musson, 1891; Smith, 1992; Stanisic et al., 2010||Including Lord Howe Island (Stanisic et al. 2010)|
|-Queensland||Present||Introduced||Not invasive||Stanisic et al., 2010||Brisbane only|
|-South Australia||Widespread||Introduced||Not invasive||Quick, 1952; Smith, 1992|
|-Tasmania||Widespread||Introduced||Not invasive||Musson, 1891; Smith, 1992|
|-Victoria||Widespread||Introduced||Not invasive||Musson, 1891; Smith, 1992|
|-Western Australia||Present||Introduced||Not invasive||Smith, 1992|
|New Zealand||Widespread||Introduced||1879||Not invasive||Quick, 1949; Barker, 1999|
History of Introduction and SpreadTop of page
There are no published records of deliberate introductions of L. maximus to areas outside its native range. It has been introduced to many countries accidentally by transport in soil, potted plants, and packaging.
Risk of IntroductionTop of page
Passive transport of L. maximus within countries is common, because of the species’ association with: soil; potted plants; stored vegetables and other produce; wooden packaging materials (boxes, crates, pellets, especially those that have been in contact with soil); and soiled agricultural and military machinery.
The species is occasionally intercepted by port quarantine personnel. Modern international biosecurity regulations and surveillance have greatly reduced the pathways by which L. maximus formerly was commonly transported across borders.
HabitatTop of page
L. maximus was probably originally a woodland and forest species, and remains common in many wooded areas of Europe (especially deciduous hornbeam-oak and beech forests). However, within both its native and adventive range L. maximus is now also widely associated with modified (anthropogenic) ecosystems, especially those with ample ground cover.
Over most of its adventive range L. maximus is certainly most prevalent in modified habitat, especially urban and rural gardens, derelict buildings, cellars, vegetable warehouses, cemeteries, parks, glasshouses and greenhouses, hedge rows, firewood stacks, and heavily vegetated waste areas such as road reserves and wooded river banks. L. maximus is common in subterranean shelters, such as cellars, military bunkers, abandoned mine shafts, municipal conduits, etc. (e.g. Dvorák, 2005; Weigand, 2014). In North America the species occurs varyingly at localities far removed from human population centres and often in little-modified habitat (e.g. Rollo and Wellington, 1975).
In reference to habitat in New Zealand, Barker (1999) observed that the species occurs along forest margins, but does not penetrate far into undisturbed temperate evergreen rainforests. It can be abundant in modified forest remnants and secondary forests (Barker, 1999, 2006). L. maximus occurs in similar situations in Chile (G.M. Barker and S. Miguel, unpublished), where forests are structurally very similar to those of New Zealand.
L. maximus penetrates subtropical to temperate rainforests in Lord Howe Island (Stanisic et al., 2010) and Hawaii (Joe and Daehler, 2008; Meyer and Cowie, 2010; Meyer et al., 2013).
Habitat ListTop of page
|Terrestrial – Managed||Cultivated / agricultural land||Secondary/tolerated habitat||Harmful (pest or invasive)|
|Cultivated / agricultural land||Secondary/tolerated habitat||Natural|
|Protected agriculture (e.g. glasshouse production)||Secondary/tolerated habitat||Harmful (pest or invasive)|
|Protected agriculture (e.g. glasshouse production)||Secondary/tolerated habitat||Natural|
|Managed forests, plantations and orchards||Principal habitat||Harmful (pest or invasive)|
|Managed forests, plantations and orchards||Principal habitat||Natural|
|Managed grasslands (grazing systems)||Secondary/tolerated habitat||Harmful (pest or invasive)|
|Managed grasslands (grazing systems)||Secondary/tolerated habitat||Natural|
|Disturbed areas||Secondary/tolerated habitat||Harmful (pest or invasive)|
|Disturbed areas||Secondary/tolerated habitat||Natural|
|Rail / roadsides||Principal habitat||Harmful (pest or invasive)|
|Rail / roadsides||Principal habitat||Natural|
|Urban / peri-urban areas||Principal habitat||Harmful (pest or invasive)|
|Urban / peri-urban areas||Principal habitat||Natural|
|Buildings||Principal habitat||Harmful (pest or invasive)|
|Terrestrial ‑ Natural / Semi-natural||Natural forests||Principal habitat||Natural|
|Natural grasslands||Secondary/tolerated habitat||Natural|
|Land caves||Secondary/tolerated habitat||Natural|
|Rocky areas / lava flows||Secondary/tolerated habitat||Natural|
|Scrub / shrublands||Principal habitat||Natural|
|Coastal dunes||Secondary/tolerated habitat||Natural|
Hosts/Species AffectedTop of page
L. maximus will feed on living plants and is capable of inflicting significant damage to garden plants. Theobald (1895) listed L. maximus as one of the three most destructive species of slug in Britain (along with Deroceras reticulatum [as Limax agreste], and Arion ater), without any justification. Taylor (1902-07) did not cite L. maximus as a pest, although he stated that it would eat young garden plants (but preferred fungi). White (1918) considered L. maximus a major pest of cultivated plants and mushrooms. For the most part the recognition of L. maximus as a pest of cultivated plants and cultivated mushrooms has diminished since the early 20th Century, in large part because it has become widely appreciated that other slug species (e.g. Deroceras reticulatum; Arion hortensis) are more pestiferous and the predominant cause of damage in the garden. Due to its large size, L. maximus is often a conspicuous member of the garden fauna and thus often erroneously assumed to be responsible for any damage observed.
There have been no quantitative studies that determine the relative contributions of L. maximus to plant losses occurring in gardens due to slugs. Nonetheless, some authors continue to mention L. maximus as a serious garden pest (e.g. Pirone, 1978; Stange, 1978; Ebeling, 2002; Kozlowski, 2012a,b; Texas Invasive Species Institute, 2004).
The greatest potential for plant damage by L. maximus in the agricultural sector is in protected cropping, such as in glasshouses and greenhouses, or where crops occur near other dense vegetation, as the high moisture conditions and availability of daylight resting sites are highly favourable to high densities and activity. Nonetheless, there are no quantitative data available to implicate L. maximus as a significant pest in these cropping situations.
In arable fields L. maximus rarely occurs at densities sufficient to present risk to crops.
Numerous cultivated plants have been recorded as being damaged by L. maximus, but the literature is clearly not comprehensive. The significance of L. maximus as a pest in commercial mushroom beds has greatly diminished with modern mushroom cultivation practices.
In non-agricultural areas, L. maximus feeds on a variety of plants (e.g. on Coincya monensis; Hipkin and Facey, 2009) and may cause plant mortality, especially in the seedling stage. For the most part this herbivory by L. maximus goes unnoticed in these situations and there is very little published information on its ecosystem-level significance. However, recent experimental work has shown that L. maximus makes a significant contribution to herbivory on seedlings in higher elevation, subtropical to temperate rainforests in Hawaii (Joe, 2006; Joe and Daehler, 2008) and in boreal forests in North America (Noel, 2004; Holloway 2008; Humber, 2009; Moss and Hermanutz, 2009, 2010; Gosse et al., 2011) which may have implications for plant recruitment in both forest systems.
It is also recognized that L. maximus feeds extensively on fungi, especially fungal fruiting structures, in woodland and forest systems (Elliott, 1922; Frömming, 1940; Keller and Snell, 2002; Halbwachs and Bässler, 2015). However, the ecosystem-level significance of this mycophagy is unknown.
Growth StagesTop of page Flowering stage, Fruiting stage, Post-harvest, Seedling stage, Vegetative growing stage
SymptomsTop of page
Damage is simply holes rasped in plant tissues. While mollusc damage is characteristic to the expert, it resembles that caused by various insects with which it is often confused.
The damage caused to plants by L. maximus is not readily differentiated from that caused by other gastropods. Even the association of L. maximus with damaged plants is not definitive evidence that the species is solely or even partially responsible.
List of Symptoms/SignsTop of page
|Fruit / external feeding|
|Fruit / frass visible|
|Growing point / external feeding|
|Growing point / frass visible|
|Inflorescence / external feeding|
|Inflorescence / frass visible|
|Leaves / external feeding|
|Leaves / frass visible|
|Leaves / shredding|
|Seeds / empty grains|
|Seeds / external feeding|
|Seeds / frass visible|
|Stems / external feeding|
|Stems / visible frass|
|Vegetative organs / external feeding|
|Vegetative organs / frass visible|
|Vegetative organs / internal feeding|
|Whole plant / external feeding|
|Whole plant / frass visible|
Biology and EcologyTop of page
Chromosome numbers n=31, 2n=62 (Garbar et al., 2011).
Morphometric analyses indicate that L. maximus is highly variable, but geographic structure in morphological variability has not been found (Garbar et al., 2011).
Reproductive Biology and Phenology
L. maximus is hermaphroditic, with individuals producing both male and female gametes. Its reproductive biology has been described by Sokolove and McCrone (1978), Barker and McGhie (1984), Wayne (2001), and others. L. maximus is a pluriennial or multivoltine, iteroparous species, with a lifespan of 2.5 to 3 years and with 2-3 egg-laying periods in their life cycle. Sokolove and McCrone (1978) and Barker and MCGhie (1984) showed that L. maximus has an annual cycle of changes in the reproductive system. Spermatogenesis was induced by the transfer from short days to long days in the spring and early summer, and this was followed by oogenesis during the summer, and fertilization followed by egg laying in the autumn. The cerebral ganglion of the central nervous system mediates the effects of long days on both male and female components of the reproductive system in transplant studies (McCrone et al., 1981). Subsequent studies demonstrated that in response to long days, both the cerebral ganglion and haemolymph contain a male gonadotropic factor that stimulates proliferation of spermatogonia in recipient slugs maintained in inhibitory short days (Melrose et al., 1983). Wayne (2001) concluded that the male gonadotropic factor probably originates in the cerebral ganglion (which contains many secretory neurons) and is secreted into the haemolymph from which it diffuses throughout the circulatory system to reach its target site(s) in the reproductive system.
L. maximus is generally considered an obligate out-crossing species following the work of McCracken and Selander (1980) and Foltz et al. (1984).
Mating in L. maximus has been described by numerous authors, including Adams (1898), Taylor (1894-1900), Kew (1901), Gerhardt (1933, 1934), Karlin and Bacon (1960), Langlois (1965), Barker and McGhie (1984), and Barker (1999). Courtship begins with circular crawling that culminates in mutual intertwining of the bodies, often on vertical surfaces such as tree trunks and walls. The slugs may then crawl over the edge of some object and become suspended by a thick thread of mucus. Mating occurs in that position, following which the slugs may decend or even drop to the ground.
Contrary to common belief, L. maximus does not produce spermatophores; mating animals simply exchange seminal material.
As in other pulmonates (Peake, 1978), there is considerable intraspecific variation in the size of L. maximus at hatching and this strongly influences subsequent development, even amongst individuals hatched from the same batch of eggs (Prior, 1983).
L. maximus is generally a solitary, ground-dwelling animal (Cook and Radford, 1988). Only when environmental conditions are highly favourable and daytime resting sites plentiful, does it occur in high local densities. L. maximus exhibits circadian rhythmicity in its locomotor activity (Sokolove et al., 1977), with onset of nocturnal foraging and other activities triggered by changes in ambient environmental conditions. Rollo (1982) developed empirical curvilinear regression and ‘limit-type’ models of regulation of activity. The models indicated the most important factors were time of day (i.e. circadian rhythm), light intensity, change in light intensity and substrate temperature, shelter temperature, length of night, time of sunset, and age and degree of hydration of the individuals. Hess and Prior (1985) found that, under dry conditions, L. maximus remained active well beyond the time that activity would have ceased under more favourable, moist conditions.
L. maximus uses daytime resting sites in sheltered microhabitats to which they repeatedly return and defend aggressively (Taylor, 1902-07; Gelperin, 1974; Rollo and Wellington, 1979; Barker and McGhie, 1984). Such use of protected daytime resting sites is most strongly displayed when daytime conditions are most unfavourable for survival of slugs in exposed microhabitats.
L. maximus does not display specific hibernatory or aestivatory behavioural or physiological adaptations, but will seek protection of sheltered microsites during periods of extreme cold, heat or dryness.
Population Size and Density
There are relatively few estimates of population densities of L. maximus in natural and modified habitats alike. Mordan (1973) and Meyer et al. (2013) recorded L. maximus at densities <1m2 in ash-oak woodland in Huntingdonshire, UK and Metrosideros rainforests in the Hawaiian Islands, respectively. This level of population density is probably typical for the species in most habitats. However, where moisture, shelter and food resources are abundant, L. maximus densities are likely to exceed 1/m2. In abandoned farm buildings in Gippsland, Victoria, Australia, for example, L. maximus populations in the order of 2-3/m2 have been recorded in night-time searches by lantern (G.M. Barker, unpubl.).
L. maximus is an omnivore, consuming fungal mycelia and fruiting structures (e.g. mushrooms), lichens, algae, fresh and decaying leaves, roots (including tubers), flowers, fruits and seeds of vascular plants, as well as domestic refuse, animal scats and faeces, prepared animal feeds such as dog biscuits and canned cat food, and both invertebrate and vertebrate carrion. The species is also cannibalistic. L. maximus will also graze biofilms on rocks, logs and tree trunks, and in buildings, the moist, mouldy wall and floor coverings. Such grazing can also extend to the covers, bindings and pages of books and other documents.
For the most part, L. maximus is considered primarily a fungivore (Webb, 1988; Taylor, 1902-07; Elliott, 1922; Frömming, 1940, 1954; Keller and Snell, 2002; Halbwachs and Bässler, 2015). Frömming (1954) found that L. maximus fed on 23 basidiomycete fungi species.
Green plants are generally considered not a major part of this species’ diet (e.g. Forsyth 2004). However, at least some populations may be largely phytophagous on higher plants (e.g. Cook and Radford, 1988) and cause damage to cultivated and wild plants alike (Barker, 1999). In an experimental situation, Frömming (1954) found that L. maximus accepted over two-thirds of offered ornamental plants and 7 of 16 lichens.
Aggressive behaviour has long been known in various slug species. Detailed accounts of the aggression in L. maximus have been given by Rollo and Wellington (1977, 1979) and by Ørmen et al. (2009, 2010). Slugs become aggressive during the late juvenile stage as they become sexually mature. Both conspecifics and other species are attacked, in some cases fatally. Rollo and Wellington (1979) found aggressiveness varied seasonally, occurring mainly during summer when hot, dry weather reduced the availability of shelter and food. Since slugs avoided shelters occupied by large aggressors and shelters closest to food were usually selected, Rollo and Wellington concluded that agonistic behaviour improved acquisition of shelter and food simultaneously.
Rollo and Wellington (1979) found mode of attack usually involved repeated biting of the victim, sometimes adopting a ‘rear-and-lunge’ strike. Barker and McGhie (1984) described how this species often inflicted severe body wounds through repeated biting.
Because it shows aggressive behaviour towards other slugs, a popular misconception that L. maximus is carnivorous and includes other slugs in its diet has arisen (see discussion in Barker, 1999; Barker and Efford, 2004). For example, because of aggression towards slugs of genus Arion, Winter et al. (2009) concluded without definitive evidence that L. maximus was probably predominantly carnivorous. This perception has crept into the ecosystem management literature (e.g. Mallick and Driessen, 2010).
ClimateTop of page
|C - Temperate/Mesothermal climate||Preferred||Average temp. of coldest month > 0°C and < 18°C, mean warmest month > 10°C|
|Cf - Warm temperate climate, wet all year||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, wet all year|
|Cs - Warm temperate climate with dry summer||Preferred||Warm average temp. > 10°C, Cold average temp. > 0°C, dry summers|
|Cw - Warm temperate climate with dry winter||Preferred||Warm temperate climate with dry winter (Warm average temp. > 10°C, Cold average temp. > 0°C, dry winters)|
|D - Continental/Microthermal climate||Preferred||Continental/Microthermal climate (Average temp. of coldest month < 0°C, mean warmest month > 10°C)|
|Df - Continental climate, wet all year||Preferred||Continental climate, wet all year (Warm average temp. > 10°C, coldest month < 0°C, wet all year)|
|Ds - Continental climate with dry summer||Preferred||Continental climate with dry summer (Warm average temp. > 10°C, coldest month < 0°C, dry summers)|
|Dw - Continental climate with dry winter||Preferred||Continental climate with dry winter (Warm average temp. > 10°C, coldest month < 0°C, dry winters)|
|ET - Tundra climate||Tolerated||Tundra climate (Average temp. of warmest month < 10°C and > 0°C)|
Natural enemiesTop of page
|Natural enemy||Type||Life stages||Specificity||References||Biological control in||Biological control on|
|Erinaceus europaeus||Predator||Adults/Juveniles||not specific||New Zealand||Gardens; horticultural crops; pastures|
|Scaphinotus striatopunctatus||Predator||Adults/Juveniles||not specific||USA (California) [experimental only]||Horticultural land|
Notes on Natural EnemiesTop of page
There is a very extensive literature on predators and parasites of land snails and slugs, but few reports specifically relate to L. maximus. The great majority of parasites and predators utilizing L. maximus are not host/prey specific.
Contrary to the common predation of birds on slugs and snails (Wild and Lawson 1937; South, 1992; Allen, 2004), Boycott (1934) considered that most bird species reject larger slugs such as L. maximus.
The ability of various species of slugs to survive attacks by predators by autotomy is well known (Stasek, 1967) and has been recorded in L. maximus (Fredj-Reygrobellet, 1975).
There have been attempts at biological control of slugs, but in very few cases has L. maximus been the target species. Altieri et al. (1982) demonstrated that a significant reduction in numbers of L. maximus could be achieved by releasing a number of adult Scaphinotus striatopunctatus (Carabidae, Cychrinae) on horticultural land.
The parasitic nematode Phasmarhabditis hermaphrodita has been developed as a biological control agent for pest slugs and snails in gardens and horticultural crops (Morand et al., 2004). However, Grewal et al. (2003a) demonstrated the inability of P. hermaphrodita to effect mortality in L. maximus and several other slug species.
In most cases the biological control agent introduced has been a generalist, and pest snails and slugs collectively have been the target. An example, is the European hedgehog (Erinaceus europaeus) introduced to New Zealand to control slugs and snails (and insect pests) in gardens, crops and pastures (Brockie, 1990). L. maximus is known to be included in the diet of the European hedgehog (Allen, 2004).
Means of Movement and DispersalTop of page
Active dispersal is commonly thought to be minimal in slugs, including L. maximus (e.g. Boycott, 1934; South, 1965), but for contrary views, see Chichester and Getz (1969). Rollo (1983b) showed L. maximus exhibited a definitive dispersal phase associated with female-phase maturation. Rollo suggested that this dispersal would be advantageous in transient field environments.
Active dispersal rates of land snails and slugs were recently reviewed by Kramarenko (2014). However, no information on L. maximus was included.
All terrestrial gastropods are dispersed with flood debris. This is most prevalent in small bodied species, or eggs and juveniles of larger species. There are no records specific to L. maximus.
Passive transport of L. maximus within countries is common, due to the species’ association with soil; potted plants; stored vegetables and other produce; wooden packaging materials (boxes, crates, pellets) (especially those that have been in contact with soil); and soiled agricultural and military machinery. The establishment of the species as an adventive in many regions of the world by the early to mid-19th Century, evidently associated with early trade and settlement by Europeans, is testament to the propensity for L. maximus to be introduced to new regions.
Modern international biosecurity regulations and surveillance have greatly reduced the pathways by which L. maximus formerly was commonly transported across borders. Nonetheless, the species is occasionally intercepted by port quarantine personnel (e.g. Hanna 1966; Dundee 1974, Roll et al. 2009; Robinson, 1999).
Robinson (1999, Table 2, p. 422) listed the principal pathways by which gastropod interceptions are made by quarantine personnel at the US border. In general terms, L. maximus would be expected to follow this pattern, with plants (horticultural), containers, cut flowers, fresh fruit, fresh vegetables and herbs, and military cargo, the most likely vectors for introduction.
Plant TradeTop of page
|Plant parts liable to carry the pest in trade/transport||Pest stages||Borne internally||Borne externally||Visibility of pest or symptoms|
|Bark||adults; eggs; juveniles||Yes||Pest or symptoms usually visible to the naked eye|
|Bulbs/Tubers/Corms/Rhizomes||adults; eggs; juveniles||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Flowers/Inflorescences/Cones/Calyx||adults; eggs; juveniles||Yes||Pest or symptoms usually visible to the naked eye|
|Fruits (inc. pods)||adults; eggs; juveniles||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Growing medium accompanying plants||adults; eggs; juveniles||Yes||Yes||Pest or symptoms usually visible to the naked eye|
|Leaves||adults; eggs; juveniles||Yes||Pest or symptoms usually visible to the naked eye|
|Seedlings/Micropropagated plants||adults; eggs; juveniles||Yes||Pest or symptoms usually visible to the naked eye|
|Stems (above ground)/Shoots/Trunks/Branches||adults; eggs; juveniles||Yes||Yes||Pest or symptoms usually visible to the naked eye|
Wood PackagingTop of page
|Wood Packaging liable to carry the pest in trade/transport||Timber type||Used as packing|
|Solid wood packing material with bark||Yes|
|Solid wood packing material without bark||Yes|
Economic ImpactTop of page
In discussing pest gastropods of Europe, Godan (1983) concluded that L. maximus “is certainly not one of the major slug pests”. This is consistent with the general view that L. maximus is not a significant plant pest. In both the native and introduced ranges, the species is however a minor pest in gardens, allotments, municipal gardens, commercial flower and vegetable crops (including protected cropping in greenhouses), cultivated mushrooms, and in stored vegetables and fruits. Occasionally, damage to plants is of sufficient intensity to warrant adoption of controls, principally the application of molluscicidal baits.
L. maximus also has indirect economic impact. The metatrongyloidean nematode parasites of ruminants, including all principal classes of farmed livestock, utilize gastropods as intermediate hosts (Anderson, 2000; Grewal et al., 2003b). These metatrongyloidean infections are of great economic significance, with loss of productivity and the need for costly antihelminthic treatments. Among these parasites are Mullerius capillaris in sheep, goats, and deer; Cystocaulus ocreatus, Neostrongylus linearis, Protostrongylus davtiani, P. hobmaieri, P. rufescens and P. skrjabini in sheep and goats; Elaphostrongylus cervi and Parelaphostrongylus tenuis in deer; and Elaphostrongylus rangiferi in reindeer. Other vertebrates of economic importance that are infected with metatrongyloidean parasites include mustelids – utilized in the fur trade, infected by Skrjabingylus nasicola, and various ruminants harvested from the wild for meat and/or fibre infected with various protostrongylid parasites. Generally these parasites are able to develop to infective stages in a great many gastropod intermediate host species. Studies to date are not exhaustive in identifying the species of intermediate hosts, but tend to focus on species locally important. That L. maximus acts as an intermediate host has been confirmed for several of these parasites (see summaries in Anderson, 2000; Grewal et al., 2003b).
Birds are also commonly infected by helminth parasites requiring gastropods as intermediate hosts. These parasites can be economically important in birds farmed or harvested from the wild for eggs, feathers and/or meat, and game birds farmed to augment wild populations. Among those parasites utilizing terrestrial gastropod hosts is the chicken tapeworm Davainea proglottina, with L. maximus among its many hosts (Graham et al. 1937; Mehlhorn and Piekarski, 1998).
Environmental ImpactTop of page
L. maximus is a distinctive native faunal element of many European wild places, including various national parks and forests. But the species is also adventive in many significant wild places internationally, including but not limited to Východné Karpaty Biosphere Reserve, Slovakia-Poland-Ukraine; Uglya Biosphere Reserve, Ukraine; Tongariro National Park and Egmont National Park, New Zealand; Tasmanian Wilderness World Heritage Area and Mount Field National Park (Tasmania), Lord Howe Island Group World Heritage Site (Lord Howe), Australia; Hakalua Forest National Wildlife Refuge and Haleakala National Park, Hawaiian Islands; Redwood National Park (California), Indiana Dunes National Lakeshore (Indiana), Glacier National Park (Montana), Black Hills National Forest (South Dakota), Olympic National Park (Washington), Mount Ranier National Park (Washington), Shenandoah National Park (Virginia) USA.; Terra Nova and Gros Morne National Parks (Newfoundland), and Ktunaxa Traditional Territory (British Columbia), Canada; Nahuel Huapi National Park, Argentina; Valdivia National Reserve, Chile; and Table Mountain National Park, South Africa. In only a few instances has there been any assessment of risk posed by L. maximus in these settings (e.g. Hutton et al., 2007; Mallick and Driessen, 2010; Meyer and Cowie, 2010; Gotthardt and Walton, 2011; Gosse et al., 2011; Walton and Gotthardt, 2011).
There are several mechanisms by which L. maximus might impact the natural environment, including herbivory and alternation of plant community structure; mycophagy, detritivory and secretory products causing alternation of nutrient cycling; predation and competitive displacement influencing gastropod community structure; and a role as an intermediate host of parasites leading to changes in host population dynamics. Nonetheless, to date, there are no documented cases of L. maximus functioning as an ecosystem engineer, where the species effects ecosystem-level changes. While documented impacts are generally considered negative (but some view L. maximus as positive in domestic gardens because of its assumed predation of pest slugs), any changes in ecosystem properties directly attributable to the species have been minor or just one small component of multiple drivers influencing the system.
L. maximus feeds on a variety of plants and may reduce plant fitness and increase mortality, especially in the seedling stage. For the most part this herbivory by L. maximus goes unnoticed and there is very little published information on its ecosystem-level significance. However, observations and experimental work has shown that L. maximus and other slugs make a contribution to herbivory on seedlings in higher elevation, subtropical to temperate rainforests in Hawaii (US Fish and Wildlife Service, 1998; Bruegmann and Caraway, 2003; US Army Garrison Hawaii, 2005; Joe, 2006; Joe and Daehler, 2008) and in boreal forests in North America (Noel, 2004; Holloway 2008; Humber, 2009; Moss and Hermanutz, 2009, 2010; Gosse et al., 2011) which, coupled with the effects of other anthropogenic disturbances (including other adventive species), may have implications for plant recruitment in both forest systems. However, to date there is no evidence that the presence of L. maximus leads to changes in vegetation communities. The studies in Hawaiian forests suggest that persistence of several rare and endangered plant species may be compromised by herbivory by L. maximus and several other adventive slugs. Among these are Alsinidendron obovatum (Caryophyllaceae) and Cyanea superba (Campanulaceae), for which threats are currently actively managed, including molluscicidal control of adventive slugs at critical sites (US Fish and Wildlife Service, 2007a,b; US Army Garrison 2009, 2010).
Macro-invertebrates play an important role in plant litter decomposition and nutrient release in forest systems (e.g. Meyer et al., 2011). In the case of terrestrial gastropods, contribution to litter decomposition occurs directly by their own metabolism and indirectly by mucus and faecal deposition enhancing micro-arthropod and microbial activity. Furthermore, species such as L. maximus are important consumers of macrofungi and probably influence decomposition and nutrient cycling through dispersal of fungal spores and hyphae. The contribution of individual species can be expected to scale with their density and biomass. L. maximus generally has low density (~ 1/m2) in forest systems, both in the native and introduced ranges, but can contribute significantly to gastropod biomass (e.g. Meyer et al., 2013). Nonetheless, there is little published information of the consequence of L. maximus colonization on nutrient cycling in forests and woods.
L. maximus is rather pugnacious. It is aggressive towards conspecifics and other slug species when competition exists over vital resources such as shelter and food (Rollo and Wellington, 1977, 1979; Barker and McGhie, 1984). Aggressiveness is more common during adverse weather conditions when the availability of shelter and food is limiting. Adults are likely to enter into interspecific competition because few refuges of suitable size tend to be available. Under experimental conditions Rollo (1983a,b) demonstrated that L. maximus can reduce the reproductive success and increase mortality of the non- or less-aggressive slug species. Busch (2007) interpreted this as indicating that L. maximus has displaced native Ariolimax columbianus in the American Pacific Northwest through competition for food resources, while Voller and McNay (2007) considered predation by L. maximus as a threat to A. columbianus. However, there is no evidence that A. columbianus has declined in range or abundance in the presence of L. maximus. Recovery plans for several endangered land snails include competition and predation by L. maximus as a potential threatening process (e.g. Parks Canada Agency, 2009; Oregon Forestsnail Recovery Team, 2012), but the risk may be overstated in the absence of sympatry and competition for food and shelter.
Threatened SpeciesTop of page
|Threatened Species||Conservation Status||Where Threatened||Mechanism||References||Notes|
|Allogona townsendiana (Oregon forestsnail)||National list(s) National list(s)||British Columbia; Oregon||Competition; Predation||Oregon Forestsnail Recovery Team, 2012|
|Alsinidendron obovatum||CR (IUCN red list: Critically endangered) CR (IUCN red list: Critically endangered)||Hawaii||Herbivory/grazing/browsing||Joe, 2006; Joe and Daehler, 2008; US Army Garrison, 2010; US Army Garrison Hawaii, 2005; US Fish and Wildlife Service, 1998|
|Cyanea superba||EW (IUCN red list: Extinct in the wild) EW (IUCN red list: Extinct in the wild)||Hawaii||Herbivory/grazing/browsing||Bruegmann and Caraway, 2003; Joe, 2006; Joe and Daehler, 2008; US Army Garrison, 2010; US Army Garrison Hawaii, 2005; US Fish and Wildlife Service, 1998|
|Hemphillia dromedarius (dromedary jumping-slug)||National list(s) National list(s)||Canada||Competition; Predation||Parks Canada Agency, 2009|
Social ImpactTop of page
In moist environments, L. maximus has a propensity to invade dwellings, feeding on biofilms and organic debris on floor and wall coverings; on books, other paper documents and wallpapers; stored vegetables and fruits; and on pet foods. This activity is generally not of economic consequence, but reduces aesthetic values.
Many gastropods are known as intermediate hosts of helminth parasites of mammals, including humans and their pets (Anderson, 2000; Grewal et al., 2003b). Angiostrongylus vasorum, also known as French heartworm, causes canine angiostrongyliasis in dogs (and various wild canines). The fox lungworm Crenosoma vulpis also occasionally parasitizes domestic dogs. L. maximus is one of a number of gastropod intermediate hosts for these parasites (Ferdushy et al., 2009). Aelurostrongylus abstrusus, the feline lungworm, and less commonly Troglostrongylus brevior and T. subcrenatus parasitise the domestic cat (and wild relatives). Although not yet confirmed, L. maximus is probably included among intermediate hosts. These parasites are normally acquired by dogs and cats after ingestion of a paratenic host such as a bird or rodent that had in turn eaten a gastropod intermediate host. These parasites are not zoonotic - that is, they cannot be communicated to humans.
Of human health significance are Angiostrongylus cantonensis and Angiostrongylus costaricensis (see Cowie, 2013; Spratt, 2015 for review), which normally infest the lungs of rats, and have a larval stage which can only live in gastropods. In humans, infection by A. cantonensis causes eosinophilic meningitis (Alicata, 1991; Cowie, 2013a), a serious condition that can lead to death or permanent brain and nerve damage. L. maximus has been implicated in the transmission of parasite in a case of eosinophilic meningitis in Sydney, Australia (Senanayake et al., 2003) and infected L. maximus have been reported in the Hawaiian Islands (Kim et al., 2014). A. costaricensis can cause severe ischemic and inflammatory intestinal lesions in humans (Cespedes et al., 1967; Graeff-Teixeira et al., 1991). Neither A. cantonensis nor A. costaricensis can complete its life cycle in human hosts. L. maximus is one of several known gastropod intermediate hosts of A. costaricensis in South America (Teixeira et al., 1993). A. cantonensis was once known to be a problem only in the tropical Indo-Pacific region, but it has since spread to other regions (Cowie, 2013a; Kim et al., 2014). Similarly, A. costaricensis infection was most prevalent in Latin America and the Caribbean but has now spread to other regions. In both cases, live gastropods (or possibly infective stage parasite larvae shed from these infected gastropods) that are accidentally eaten with improperly cleaned vegetables (especially salads), or gastropods that are deliberatly eaten raw or that have been improperly cooked, can act as vectors for the parasite (Cowie, 2013b).
Risk and Impact FactorsTop of page Invasiveness
- Has a broad native range
- Abundant in its native range
- Is a habitat generalist
- Tolerates, or benefits from, cultivation, browsing pressure, mutilation, fire etc
- Pioneering in disturbed areas
- Tolerant of shade
- Capable of securing and ingesting a wide range of food
- Benefits from human association (i.e. it is a human commensal)
- Long lived
- Host damage
- Infrastructure damage
- Modification of successional patterns
- Negatively impacts agriculture
- Negatively impacts human health
- Negatively impacts animal health
- Threat to/ loss of endangered species
- Damages animal/plant products
- Interaction with other invasive species
- Highly likely to be transported internationally accidentally
- Difficult/costly to control
Detection and InspectionTop of page
Detection of L. maximus would usually involve searches for the animals, either as eggs or slugs, among leaf and woody debris on the ground. Active L. maximus may be detected by searches made at night with the aid of a lantern. Such searches can be augmented by trapping methods, typically boards, tiles or wet jute sacks laid on the ground and inspected regularly for presence of slugs sheltering underneath. The ‘catch’ of slugs can be improved in some cases by placing some molluscicide baits under the trap. An alternative trapping method relies on open vessels containing an attractant or preservative placed on the ground, and likewise examined periodically to presence of slugs. Attractants include beer, milk, cat food, or milled grains.
In leafy crops such as lettuce and cabbage, individual plants should be inspected for presence of slugs within or under the plants.
Damage to plants should not be attributed to L. maximus in the absence of confirmed feeding by this species on the plants in question as other gastropod species may be responsible. The definitive proof of culpability is observation of L. maximus actually feeding on the plants. This will generally require search of the crop at night with the aid of a lantern.
In shipped commodities, detection of L. maximus generally will require manual searches, although in large consignments such as shipping containers, there is the option of augmenting these searches with open vessel traps as described above. Often the presence of L. maximus, and other gastropods, may be indicated by the mucus trails left by the active animals.
Similarities to Other Species/ConditionsTop of page
L. maximus is highly variable in body coloration. Numerous variety names are to be found in the literature - see Taylor (1902-07), Hesse (1926) and Quick (1949, 1960).
L. maximus has frequently been mistaken for L. cinereoniger, but is distinguished externally from that species by its shorter, less prominent keel; smaller tubercles on the body; uniformly pale sole; its banding or maculation; absence of a pale stripe running from keel to posterior margin of mantle; and uniformly vinous brown tentacles. Internally L. maximus is distinguished by its larger shell; larger jaw; the general weakness of the ectocones and endocones on the radular teeth; and shorter phallus (relative to body length) in the reproductive system. Juveniles of L. cinereoniger may be confused with adult L. maximus due to their uniformly colored sole.
L. maximus is externally similar to several Limax species (e.g. Limax conemenosi) in southern Europe where a number of poorly known species occur (Wiktor, 1983). Dissection is usually needed to determine specific status.
L. maximus juveniles may sometimes be confused with Lehmannia species (Limacidae). However, unlike several Lehmannia species the lateral bands never occur on the mantle.
Prevention and ControlTop of page
Due to the variable regulations around (de)registration of pesticides, your national list of registered pesticides or relevant authority should be consulted to determine which products are legally allowed for use in your country when considering chemical control. Pesticides should always be used in a lawful manner, consistent with the product's label.
As with other terrestrial gastropod species, once established in an area, eradication is most unlikely. The most effective management option includes a combination of prevention of spread, early detection and rapid response, and control. To prevent species such as L. maximus from spreading, public education can raise awareness to reduce human mediated spread, coupled with robust border inspection and quarantine procedures. Early detection and rapid response would strategically involve surveillance by biologists, and a platform to enable the public to report sightings. Control might involve physically removing slugs and their eggs, but would generally rely on chemical treatments (principally baits containing molluscicides).
Eradication is considered unachievable except perhaps in the earliest stages of incursion.
Eradication of terrestrial gastropods is generally not attempted, mainly because most infested areas are surrounded by habitat occupied by the species and therefore subject to recolonisation by immigration. It is also very difficult to achieve 100% mortality because of differential susceptibility of individuals in populations and, in the case of chemical or manual controls, there are no barriers to rapid population resurgence from survivors, hatch from eggs, and recolonisation by immigration. Thirdly, most plant protection control operations are concerned only with short-term release of plants (usually seedlings) from herbivory. Finally, current biological and chemical molluscicides affect both target and sympatric non-target mollusc species.
The eradication of L. maximus will be impossible unless the populations are naturally isolated or barriers to recolonisation (re-invasion) are established as part of the management regime. In isolated populations, eradication is theoretically possible by repeated molluscicide application over several years (reproductive seasons). Experience with other gastropod pest species clearly demonstrates that it is feasible to eradicate incipient infestations, but to date there have been no successful attempts to eradicate long-established populations of invasive gastropod species.
Cultural control and sanitary measures
Hand collection with subsequent destruction of animals is the oldest method of control of pest gastropods (Godan, 1983), and has been used effectively in conjunction with chemical methods for management of infestations in agricultural areas and in eradication of incipient infestations of invasive species.
However, manual removal as a control method has three primary constraints: the difficulty of finding sufficient numbers of individuals to effectively manage or eradicate the population; the high labour costs; and the physical disturbance to the habitat. These constraints are highest where the species/life stages being controlled are small and therefore difficult to detect in infested habitat. Adults of L. maximus are large and thus potentially amenable to manual collection and destruction. However, the eggs and juveniles are much smaller and often concealed in soil and under and within plant debris, reducing the prospects for control by manual removal.
The abundance and damage potential of L. maximus and other gastropod pests in garden and agricultural situations is often closely linked to the amount of ground cover, as refuge for the pests, adjacent to the cultivated plants. Where growing areas are surrounded with grass or weedy headlands, slug activity and damage is generally higher than where the surrounds are free of ground cover plants. In gardens and horticultural crops, risk is heightened when there is an accumulation of debris such as boards, pots, cardboard and weed growth.
There has been much interest in biological control approaches to management of pest gastropods. A large number of natural enemies of terrestrial gastropods are known, with a voluminous, albeit widely dispersed and often specialized, literature. Recorded enemies include: pathogenic bacteria, fungi, and viruses; parasitic protozoa, ciliophora, parabasalia microsporidia, nematodes, trematodes and cestods; parasitoid and predatory Sciomyzidae; predatory gastropods, flatworms and arthropods; and predatory vertebrates, including amphibians, reptiles, birds, and mammals. Some of these records relate to L. maximus. Most natural enemies of terrestrial gastropods have proved not to be host-specific and therefore are not amenable to use in control programmes where effects on non-target species are of concern (Cowie, 2001). To date, no natural enemy specific to L. maximus is known.
There has been recent interest in inundative biological control approaches to control of terrestrial gastropod pests. These inundative approaches have primarily focused on the rhabiditid nematode Phasmarhabditis hermaphrodita (Morand et al. 2004), with the development of a commercial product. This species is native to Europe. Its use in agriculture there is at present constrained by the high cost. The lack of host-specificity largely negates the potential use of P. hermaphrodita outside its native European range, although the discovery of the nematode in other parts of the world, as an adventive, continues to stimulate commercial interests. Nonetheless, P. hermaphrodita is ineffective as a control agent for L. maximus (Grewal et al., 2003a).
Application of pesticides (molluscicides) is regarded as the most pragmatic approach to control of terrestrial gastropod pests. Delivery of molluscicidal chemicals to target pest populations has primarily focused on bait formulations, and there has been substantial investment in bait technologies by government and commercial agencies in many parts of the world. In general, other molluscicide formulations, such as sprays and dusts, have proved ineffective in control of field infestations. There are considerable biological imperatives for this, not least the demonstrated remarkably difficult delivery of pesticides to the target gastropod tissues when applied as spray and dust formulations. The mucus extruded on the body surface provides a significant barrier to uptake of pesticides of large molecular weight by gastropods active in treated environments. Both sprays and dusts as means of control depend on high application rates of active ingredient and are more indiscriminate in their effects on non-target fauna than bait formulations.
There are three major classes of compounds presently used in control of terrestrial gastropod pests, namely metaldehyde, carbamates, and metal chelates (see discussion by Barker and Watts, 2002).
Under the relatively uniform conditions of agricultural fields, the level of control of pest gastropods with a single baiting operation is rarely above 70%, and typically 10–60% (e.g. Frömming and Plate, 1952; Godan 1983; Barker et al., 1991). The level of control in more complex, spatially varied ecosystems can be expected to be substantially less. As discussed by Barker and Watts (2002), the efficacy of molluscicide bait treatments is influenced by a number of factors, including: variation in susceptibility within species with age, size, and reproductive condition; variation in effectiveness with bait formulation; dependency of toxicity on dose and environment; dependency of level of control on placement of baits at sites of gastropod activity.
Most commercial bait formulations for slug and snail control are known to be attractive, and the mollucicidal ingredients toxic to L. maximus. Nonetheless, juveniles of L. maximus are considered more tolerant of molluscides than are adults (Godan, 1983).
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08/03/2015 Original text by:
Dr Gary Barker, Landcare Research, Private Bag 3127, Hamilton, New Zealand
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