UNITED STATES

DEPARTMENT OF COMMERCE

Penny Pritzker

Secretary

NATIONAL OCEANIC AND

ATMOSPHERIC ADMINISTRATION

Kathy Sullivan

Acting, Under Secretary for Oceans & Atmosphere

NOAA Administrator

NAT OI NAL OCEANIC AND ATMOSPHERIC ADMINISTRAT OI N

U.S.DEPARTMENT OF COMMERCE

NOAA Technical Memorandum GLERL-168

An overview of the management of established nonindigenous species in the 

Great Lakes

Rochelle Sturtevant1

Lauren Berent2

Thomas Makled2

Whitney Conard2

Abigail Fusaro3

Edward Rutherford4

1NOAA Sea Grant, 4840 S. State Rd., Ann Arbor, MI 48108

2Cooperative Institute for Limnology and Ecosystems Research, 4840 S. State Rd., Ann Arbor, MI 48108

3Wayne State University, 42 W. Warren Ave., Detroit, MI 48202

4NOAA Great Lakes Environmental Research Laboratory, 4840 S. State Rd., Ann Arbor, MI 48108

February 2016
ii

NOTICE

Mention of a commercial company or product does not constitute an endorsement by the NOAA. 

Use of information from this publication concerning proprietary products or the tests of such 

products for publicity or advertising purposes is not authorized. This is GLERL Contribution No. 

1795.

This publication is available as a PDF file and can be downloaded from GLERL’s web site: 

www.glerl.noaa.gov. Hard copies can be requested from GLERL Information Services, 

4840 S. State Rd., Ann Arbor, MI 48108.

pubs.glerl@noaa.gov.

NOAA’s Mission – To understand and predict changes in Earth’s environment and conserve and 

manage coastal and marine resources to meet our nation’s economic, social, and environmental 

needs 

NOAA’s Mission Goals:

• Protect, restore and manage the use of coastal and ocean resources through an ecosystem

approach to management

• Understand climate variability and change to enhance society’s ability to plan and respond

• Serve society’s needs for weather and water information

• Support the Nation’s commerce with information for safe, efficient, and environmentally

sound transportation

• Provide critical support for NOAA’s Mission
1

TABLE OF CONTENTS

1. Introduction........................................................................................................................ 11

2. Methods ............................................................................................................................. 12

3. Nonindigenous Species Management................................................................................ 16

3.1 Regulations Targeting Pathways .....................................................................................................17

3.2 Federal Regulations Targeting Specific Species...............................................................................17

3.3 State Regulations Targeting Specific Species ..................................................................................18

3.3.1 Fish pathogens.................................................................................................................... 18

3.3.2 Fish...................................................................................................................................... 21

3.3.3 Mollusks.............................................................................................................................. 26

3.3.4 Waterfleas and Mysids....................................................................................................... 27

3.3.5 Plants.................................................................................................................................. 29

3.3.6 Miscellaneous taxa covered only under the whitelist approach – Algae and Aquatic 

Invertebrates................................................................................................................................. 33

3.4 Unregulated Species........................................................................................................................34

4. Nonindigenous Species Control ......................................................................................... 35

4.1 Algae................................................................................................................................................35

4.2 Plants...............................................................................................................................................36

4.3 Fish...................................................................................................................................................56

4.4 Mollusks...........................................................................................................................................62

4.5 Insects..............................................................................................................................................65

4.6 Free-Living Crustaceans, including Waterfleas, Copepods, Mysids and Gammarids......................65

4.7 Free-Living Worms including Annelids and Playthelminthes..........................................................69

4.8 Bryozoans, Hydrozoans and Testate Amoebae...............................................................................70

4.9 Parasites and Diseases.....................................................................................................................71
2 

4.9.1 Parasitic Platyhelminthes ................................................................................................... 71

4.9.2 Parasitic copepods .............................................................................................................. 72

4.9.3 Protozoan Parasites ............................................................................................................ 73

4.9.4 Bacteria ............................................................................................................................... 75

4.9.5 Viruses ................................................................................................................................ 77 

5. Discussion ........................................................................................................................... 78

6. Literature Cited ................................................................................................................... 80

7. Acknowledgements .......................................................................................................... 129

Appendix A. Species Management Profiles ................................................................................ 131

A.1 Algae ...................................................................................................................................................131

Actinocyclus normanii f. subsalsa (Juhlin-Dannfelt) Hustedt, 1957 .......................................... 131

Bangia atropurpurea (Roth) Agardh, 1824 ................................................................................ 131

Chaetoceros muelleri subsalsum J. R. Johansen and Rushforth, 1985 ...................................... 132

Chroodactylon ornatum (C. Agardh) Basson, 1979 .................................................................... 132

Contricribra guillardii (Hasle) K. Stachura-Suchoples & D.M. Williams ..................................... 132

Cyclotella atomus Hustedt, 1937 ............................................................................................... 132

Cyclotella cryptica Reimann, Lewin, and Guillard, 1963 ............................................................ 133

Cylindrospermopsis raciborskii (Wolosz.) Seenayya and Subbaraju, 1972 ............................... 133

Diatoma ehrenbergii Kützing, 1844 ........................................................................................... 134

Discostella pseudostelligera (Hustedt) Houk and Klee, 1939 .................................................... 134

Discostella woltereckii Hustedt, 1942 ........................................................................................ 135

Hymenomonas roseola Stein, 1878 ........................................................................................... 135

Nitellopsis obtusa (Desvaux in Loiseleur) J. Groves, (1919) ....................................................... 135

Pleurosira laevis (Ehrenberg) Compère, (1843) 1982 ................................................................ 135
3

Skeletonema potamos (Weber) Hasle in Hasle & Evensen, (1970)........................................... 136

Skeletonema subsalsum (Cleve-Euler) Bethge, (1912) 1928 ..................................................... 136

Sphacelaria fluviatilis Jao, 1943................................................................................................. 136

Sphacelaria lacustris Schloesser and Blum, 1980...................................................................... 136

Stephanodiscus binderanus Krieger, 1927................................................................................. 136

Stephanodiscus subtilis (Van Goor) A. Cleve, 1951.................................................................... 137

Thalassiosira baltica Ostenfeld, 1901........................................................................................ 137

Thalassiosira lacustris (Grunow) Hasle in Hasle and Fryxell, 1977............................................ 137

Thalassiosira pseudonana (Hustedt) Hasle and Heimdal, (1957) 1970..................................... 138

Thalassiosira weissflogii (Grunow) G. Fryxell & Hasle, (1896) 1977 ......................................... 138

Ulva (Enteromorpha) flexuosa subsp. flexuosa and flexuosa subsp. paradoxa (Wolfen ex Roth) 

J. Agardh, 1883............................................................................................................................ 138

Ulva (Enteromorpha) intestinalis Linnaeus, 1753 ..................................................................... 139

Ulva (Enteromorpha) prolifera O.F. Müller, 1778 ..................................................................... 139

A.2 Annelids..............................................................................................................................................139

Branchiura sowerbyi .................................................................................................................. 139

Gianius aquaedulcis ................................................................................................................... 140

Potamothrix bedoti (Piquet, 1913) ............................................................................................ 141

Potamothrix moldaviensis Vejdovsky and Mrazek, 1902.......................................................... 141

Potamothrix vejdovskyi Hrabe, 1941......................................................................................... 142

Ripistes parasita......................................................................................................................... 143

A.3 Bacteria...............................................................................................................................................143

Aeromonas salmonicida Emmerich and Weible, 1890.............................................................. 143

Piscirickettsia cf. salmonis ......................................................................................................... 146

Renibacterium (Corynebacterium) salmoninarum Sanders and Fryer, 1980 ........................... 146
4

A.4 Bryozoa ...............................................................................................................................................148

Lophopodella carteri (Hyatt, 1865)............................................................................................ 148

A.5 Coelenterates......................................................................................................................................149

Cordylophora caspia (Pallas, 1771)............................................................................................ 149

Craspedacusta sowerbyi Lankester, 1880.................................................................................. 149

A.6 Amphipods..........................................................................................................................................150

Echinogammarus ischnus (Stebbing, 1899)............................................................................... 150

Gammarus tigrinus Sexton 1939................................................................................................ 151

A.7 Cladocerans ........................................................................................................................................151

Bosmina coregoni Baird, 1857 ................................................................................................... 151

Bythotrephes longimanus.......................................................................................................... 152

Cercopagis pengoi ...................................................................................................................... 153

Daphnia galeata galeata Sars, 1864 ......................................................................................... 155

Daphnia lumholtzi ...................................................................................................................... 156

Eubosmina maritima P.E. Müller, 1867 ..................................................................................... 156

A.8 Copepods............................................................................................................................................157

Argulus japonicus Thiele, 1900 .................................................................................................. 157

Cyclops strenuus Fischer, 1851................................................................................................... 157

Eurytemora affinis Poppe, 1880 ................................................................................................ 158

Heteropsyllus nr. nunni Coull..................................................................................................... 159

Megacyclops viridis Jurine, 1820 ............................................................................................... 159

Neoergasilus japonicus Harada, 1930........................................................................................ 160

Nitokra hibernica Brady, 1880 ................................................................................................... 160

Nitokra incerta Richard, 1893 .................................................................................................... 161
5

Salmincola lotae Olsson, 1877 ................................................................................................... 161

Schizopera borutzkyi Monchenko, 1967.................................................................................... 162

Skistodiaptomus pallidus Herrick, 1879 .................................................................................... 163

A.9 Mysids.................................................................................................................................................163

Hemimysis anomala G.O. Sars, 1907 ......................................................................................... 163

A.10 Fishes................................................................................................................................................164

Alosa aestivalis Mitchill, 1814.................................................................................................... 164

Alosa pseudoharengus Wilson, 1811......................................................................................... 165

Apeltes quadracus Mitchill, 1815............................................................................................... 166

Carassius auratus Linnaeus, 1758.............................................................................................. 167

Cyprinus carpio Linnaeus, 1758.................................................................................................. 169

Enneacanthus gloriosus Holbrook, 1855 ................................................................................... 170

Esox niger Lesueur, 1818............................................................................................................ 172

Gambusia affinis Baird and Girard, 1853................................................................................... 173

Gymnocephalus cernua Linnaeus, 1758 .................................................................................... 173

Ictiobus cyprinellus Valenciennes in Cuvier and Valenciennes, 1844........................................ 176

Lepisosteus platostomus Rafinesque, 1820............................................................................... 176

Lepomis microlophus Günther, 1859 ......................................................................................... 177

Misgurnus anguillicaudatus Cantor, 1842................................................................................. 178

Morone americana Gmelin, 1789 .............................................................................................. 179

Neogobius melanostomus Pallas, 1814 ..................................................................................... 180

Notropis buchanani Meek, 1896................................................................................................ 181

Oncorhynchus gorbuscha Walbaum, 1792 ................................................................................ 182

Oncorhynchus kisutch Walbaum, 1792...................................................................................... 184
6

Oncorhynchus mykiss Walbaum, 1792 ...................................................................................... 186

Oncorhynchus nerka (Walbaum in Artedi, 1792)....................................................................... 188

Oncorhynchus tshawytscha Walbaum in Artedi, 1792.............................................................. 189

Osmerus mordax Mitchill, 1814 ................................................................................................. 191

Petromyzon marinus (Linnaeus, 1758) ...................................................................................... 193

Phenacobius mirabilis (Girard, 1856)......................................................................................... 195

Proterorhinus semilunaris (Heckel, 1837).................................................................................. 195

Salmo trutta Linnaeus, 1758 ...................................................................................................... 197

Scardinius erythrophthalmus Linnaeus, 1758 ........................................................................... 199

A.11 Insects...............................................................................................................................................200

Acentria ephemerella Olivier, 1791 ........................................................................................... 200

Tanysphyrus lemnae Paykull/Fabricius, 1792............................................................................ 200

A.12 Mollusks............................................................................................................................................201

Corbicula fluminea O. F. Müller, 1774 ....................................................................................... 201

Dreissena polymorpha Pallas, 1771 ........................................................................................... 202

Dreissena rostriformis bugensis Andrusov, 1897 ...................................................................... 203

Lasmigona subviridis Conrad, 1835 ........................................................................................... 205

Pisidium amnicum Müller, 1774 ................................................................................................ 206

Pisidium henslowanum Shepard, 1825...................................................................................... 206

Pisidium moitessierianum Paladilhe, 1866 ................................................................................ 207

Pisidium supinum Schmidt 1850 ................................................................................................ 207

Sphaerium corneum Linnaeus, 1758.......................................................................................... 208

Bithynia tentaculata................................................................................................................... 209

Cipangopaludina chinensis malleata......................................................................................... 209
7

Cipangopaludina japonica ......................................................................................................... 210

Elimia virginica ........................................................................................................................... 210

Gillia altilis.................................................................................................................................. 211

Potamopyrgus antipodarum...................................................................................................... 211

Radix auricularia ........................................................................................................................ 212

Valvata piscinalis........................................................................................................................ 213

Viviparus georgianus I. Lea, 1834 .............................................................................................. 213

A.13 Plants................................................................................................................................................214

Agrostis gigantea Roth............................................................................................................... 214

Alnus glutinosa (L.) Gaertn......................................................................................................... 215

Alopecurus geniculatus L. .......................................................................................................... 215

Butomus umbellatus L................................................................................................................ 216

Cabomba caroliniana ................................................................................................................. 217

Carex acutiformis Ehrh............................................................................................................... 219

Carex disticha Huds. ................................................................................................................... 220

Chenopodium glaucum .............................................................................................................. 220

Cirsium palustre.......................................................................................................................... 220

Conium maculatum .................................................................................................................... 222

Echinochloa crus-galli L. P. Beauvois.......................................................................................... 224

Epilobium hirsutum L.................................................................................................................. 226

Frangula alnus P. Mill................................................................................................................. 228

Glyceria maxima......................................................................................................................... 230

Hydrocharis morsus-ranae L. ..................................................................................................... 231

Impatiens glandulifera............................................................................................................... 232
8

Iris pseudacorus.......................................................................................................................... 233

Juncus compressus Jacq.............................................................................................................. 235

Juncus gerardii Loisel.................................................................................................................. 235

Juncus inflexus L. ........................................................................................................................ 236

Lupinus polyphyllus.................................................................................................................... 237

Lycopus asper ............................................................................................................................. 237

Lycopus europaeus L. ................................................................................................................. 237

Lysimachia nummularia............................................................................................................. 238

Lysimachia vulgaris.................................................................................................................... 238

Lythrum salicaria........................................................................................................................ 239

Marsilea quadrifolia................................................................................................................... 241

Mentha aquatica L. .................................................................................................................... 242

Mentha gracilis Sole (pro sp.) .................................................................................................... 242

Mentha spicata........................................................................................................................... 243

Myosotis scorpioides L. .............................................................................................................. 243

Myosoton aquaticum (L.) Moench............................................................................................. 243

Myriophyllum spicatum L........................................................................................................... 244

Najas marina L............................................................................................................................ 246

Najas minor All. .......................................................................................................................... 247

Nasturtium officinale ................................................................................................................. 247

Nymphoides peltata................................................................................................................... 248

Pluchea odorata succulenta (Fern.) Cronq. ............................................................................... 249

Pluchea odorata odorata (L.) Cass............................................................................................. 249

Poa trivialis L. ............................................................................................................................. 249
9

Polygonum persicaria L.............................................................................................................. 250

Potamogeton crispus ................................................................................................................. 251

Puccinellia distans (Jacq.) Parl.................................................................................................... 252

Rorippa sylvestris (L.) Bess. ........................................................................................................ 252

Rumex longifolius DC. ................................................................................................................ 253

Rumex obtusifolius..................................................................................................................... 254

Salix alba..................................................................................................................................... 255

Salix fragilis ................................................................................................................................ 256

Salix purpurea............................................................................................................................. 257

Solanum dulcamara ................................................................................................................... 258

Solidago sempervirens L. ........................................................................................................... 258

Sparganium glomeratum (Laestad.) L. Neum............................................................................ 259

Trapa natans .............................................................................................................................. 259

Typha angustifolia L................................................................................................................... 260

Veronica beccabunga................................................................................................................. 261

A.14 Platyhelminthes................................................................................................................................262

Bothriocephalus acheilognathi Yamaguti, 1934........................................................................ 262

Dactylogyrus amphibothrium Wagener or Wegener, 1857...................................................... 262

Dactylogyrus hemiamphibothrium Ergens, 1956...................................................................... 263

Dugesia polychroa Schmidt, 1861.............................................................................................. 264

Ichthyocotylurus pileatus........................................................................................................... 264

Neascus brevicaudatus von Nordmann, 1832 ........................................................................... 265

Scolex pleuronectis Müller, 1788 ............................................................................................... 265

Timoniella sp. ............................................................................................................................. 265
10

A.15 Protozoa............................................................................................................................................265

Acineta nitocrae ......................................................................................................................... 265

Glugea hertwigi Weissenberg, 1911.......................................................................................... 265

Heterosporis sp........................................................................................................................... 265

Myxobolus cerebralis Hofer, 1903 ............................................................................................. 266

Psammonobiotus communis...................................................................................................... 269

Psammonobiotus dziwnowi....................................................................................................... 269

Psammonobiotus linearis........................................................................................................... 269

Sphaeromyxa sevastopoli .......................................................................................................... 269

Trypanosoma acerinae Brumpt, 1906 ....................................................................................... 269

A.16 Viruses ..............................................................................................................................................270

Novirhabdovirus sp. genotype IV sublineage b ......................................................................... 270

Ranavirus.................................................................................................................................... 272

Rhabdovirus carpio .................................................................................................................... 272
11

AN OVERVIEW OF THE MANAGEMENT OF ESTABLISHED NONINDIGENOUS 

SPECIES IN THE GREAT LAKES

Rochelle Sturtevant, Lauren Berent, Thomas Makled, Abigail Fusaro, and Ed Rutherford

1. Introduction

The Great Lakes are host to thousands of native fishes, invertebrates, plants, and other species 

that not only provide recreational and economic value to the region, but also hold important 

ecological value. However, with over 180 documented aquatic nonindigenous species1(ANS) 

and an apparent introduction rate estimated at 1.3-1.8 species·year-1through 2006, the Great 

Lakes basin is considered one of the most heavily invaded aquatic systems in the world (Mills et 

al. 1993, Ricciardi 2006, GLRI Task Force 2010). Some of these nonindigenous species may 

become invasive (i.e. “species whose introduction does or is likely to cause economic or 

environmental harm or harm to human health” (Executive Order 13112, 1999)) and threaten the 

ecological and/or socio-economic value of the Great Lakes. In contrast, other nonindigenous 

species are capable of contributing value to the Great Lakes. Pacific salmonids, for instance, are 

stocked annually by the millions to help support the Great Lakes’ multi-billion dollar fishery 

(Kocik and Jones 1999, USFWS/GLFC 2010, USACE 2012a).

This study provides a snapshot of management of established nonindigenous species in the Great 

Lakes region as a benchmark for improving management practices. We review the relevant state 

and federal regulations that impact AIS management as well as management alternatives in a 

format that allows cross-jurisdictional and cross-taxa analysis. This effort is part of a larger 

project funded by the Great Lakes Restoration Initiative to enhance the Great Lakes Aquatic 

Nonindigenous Species Information System (GLANSIS1), an online database containing 

information on the identification, distribution, ecology, impact, and management of all 

established ANS in the Great Lakes. Previously, we assessed the relative ecological and 

socioeconomic impacts of nonindigenous species (NOAA Technical Memorandum 161 – An 

Impact Assessment of Great Lakes Aquatic Nonindigenous Species; Sturtevant et al. 2014). 

 

1

These nonindigenous aquatic species have populations established in the Great Lakes basin below the ordinary 

high water mark, including connecting channels, wetlands, and waters ordinarily attached to the lakes (see 

definitions and criteria for listing in the Great Lakes Aquatic Nonindigenous Species Information System at 

http://www.glerl.noaa.gov/res/Programs/glansis/glansis.html).
12

2. Methods

This document is a synthesis of management data gathered for GLANSIS over the period 2010-

2014. GLANSIS fact sheets were primarily researched and written by the authors of this 

manuscript with some additional support from students mentioned in the Acknowledgements. 

The 181 species-specific fact sheets, which include information on ecology, invasion history,

impact, as well as management were written independently to a common template, but each was 

reviewed by the core management team for consistency as well as sent to external taxonomic 

experts (also listed in Acknowledgements) for review prior to posting in GLANSIS. The 

management sections of each fact sheet were extracted from the GLANSIS database and 

collectively form the ‘raw’ dataset (included in Appendix A), which is analyzed and summarized 

in this memo. 

Table 1. Great Lakes Established Nonindigenous Species included in GLANSIS

Scientific Name Common Name

Algae

Actinocyclus normanii f. subsalsa Diatom

Bangia atropurpurea Red Alga

Chaetoceros muelleri Diatom

Chroodactylon ornatum Red Alga

Contracribra guillardii Diatom

Cyclotella atomus Diatom

Cyclotella cryptica Diatom

Cylindrospermopsis raciborskii Cylindro

Diatoma ehrenbergii Diatom

Discostella pseudostelligera Diatom

Discostella woltereckii Diatom

Hymenomonas roseola Coccolithophorid

Nitellopsis obtusa Starry Stonewort

Pleurosira laevis Diatom

Skeletonema potamos Diatom

Skeletonema subsalsum Diatom

Sphacelaria fluviatilis Brown Alga

Sphacelaria lacustris Brown Alga

Stephanodiscus binderanus Diatom

Stephanodiscus subtilis Diatom

Thalassiosira baltica Diatom

Thalassiosira lacustris Diatom

Thalassiosira pseudonana Diatom

Thalassiosira weissflogii Diatom

Ulva flexuosa Grass Kelp

Ulva intestinalis Grass Kelp

Ulva prolifera Sea Lettuce
13

Annelids

Branchiura sowerbyi Tubificid Worm

Gianius aquaedulcis Tubificid Worm

Potamothrix bedoti Tubificid Worm

Potamothrix moldaviensis Tubificid Worm

Potamothrix vejdovskyi Tubificid Worm

Ripistes parasita Oligochaete

Bacteria

Aeromonas salmonicida Furunculosis

Piscirickettsia cf. salmonis Muskie Pox

Renibacterium salmoninarum Bacterial Kidney Disease (BKD)

Bryozoa

Lophopodella carteri Freshwater Byrozoan

Coelenterates

Cordylophora caspia Freshwater Hydroid

Craspedacusta sowerbyi Freshwater Jellyfish

Amphipods

Echinogammarus ischnus Scud

Gammarus tigrinus Amphipod

Cladocerans

Bosmina coregoni Waterflea

Bythotrephes longimanus Spiny Waterflea

Cercopagis pengoi Fishhook Waterflea

Daphnia galeata galeata Waterflea

Daphnia lumholtzi Waterflea

Eubosmina maritima Cladoceran

Copepods

Argulus japonicus Japanese Fishlouse

Cyclops strenuus Copepod

Eurytemora affinis Calanoid Copepod

Heteropsyllus nr. nunni Harpacticoid Copepod

Megacyclops viridis Cyclopoid Copepod

Neoergasilus japonicus Parasitic Copepod

Nitokra hibernica Harpacticoid Copepod

Nitokra incerta Harpacticoid Copepod

Salmincola lotae Parasitic Copepod

Schizopera borutzkyi Oarsman

Skistodiaptomus pallidus Calanoid Copepod

Mysids

Hemimysis anomala Bloody Red Shrimp

Fish

Alosa aestivalis Blueback Herring

Alosa pseudoharengus Alewife

Apeltes quadracus Fourspine Stickleback

Carassius auratus Goldfish
14

Cyprinus carpio Common Carp

Enneacanthus gloriosus Bluespotted Sunfish

Esox niger Chain Pickerel

Gambusia affinis Western Mosquitofish

Gymnocephalus cernua Ruffe

Ictiobus cyprinellus Bigmouth Buffalo

Lepisosteus platostomus Shortnose Gar

Lepomis microlophus Redear Sunfish

Misgurnus anguillicaudatus Oriental Weatherfish

Morone americana White Perch

Neogobius melanostomus Round Goby

Notropis buchanani Ghost Shiner

Oncorhynchus gorbuscha Pink Salmon

Oncorhynchus kisutch Coho Salmon

Oncorhynchus mykiss Rainbow Trout

Oncorhynchus nerka Sockeye Salmon

Oncorhynchus tshawytscha Chinook Salmon

Osmerus mordax Rainbow Smelt

Petromyzon marinus Sea Lamprey

Phencobius mirabilis Suckermouth Minnow

Proterorhinus semilunaris Tubenose Goby

Salmo trutta Brown Trout

Scardinius erythrophthalmus Rudd

Insects

Acentria ephemerella European Water Moth

Tanysphyrus lemnae Duckweed Weevil

Mollusks

Corbicula fluminea Asian Clam

Dreissena polymorpha Zebra Mussel

Dreissena rostriformis bugensis Quagga Mussel

Lasmigona subviridis Green Floater

Pisidium amnicum Greater European Peaclam

Pisidium henslowanum Henslow’s Peaclam

Pisidium moitessierianum Pygmy Peaclam

Pisidium supinum Humpbacked Peaclam

Sphaerium corneum European Fingernail Clam

Bithynia tentaculata Mud Bithynia, Faucet Snail

Cipangopaludina chinensis malleata Chinese Mysterysnail

Cipangopaludina japonica Japanese Mysterysnail

Elimia virginica Piedmont Elimia

Gillia altilis Buffalo Pebblesnail

Potamopyrgus antipodarum New Zealand Mudsnail

Radix auricularia European Ear Snail

Valvata piscinalis European Stream Valvata

Viviparus georgianus Banded Mysterysnail
15

Plants

Agrostis gigantea Redtop, Black Bent, Water Bentgrass

Alnus glutinosa Black Alder

Alopecurus geniculatus Water Foxtail, Marsh Meadow-Foxtail

Butomus umbellatus Flowering Rush

Cabomba caroliniana Carolina Fanwort

Carex acutiformis Swamp Sedge

Carex disticha Tworank Sedge

Chenopodium glaucum Oak-leaved Goosefoot

Cirsium palustre Marsh Thistle

Conium maculatum Poison Hemlock

Echinochloa crus-galli Barnyard Grass

Epilobium hirsutum Great Hairy Willow Herb

Frangula alnus Glossy Buckthorn

Glyceria maxima Reed Mannagrass

Hydrocharis morsus-ranae European Frogbit

Impatiens glandulifera Ornamental Jewelweed

Iris pseudacorus Yellow Iris

Juncus compressus Flattened Rush

Juncus gerardii Black-grass Rush

Juncus inflexus European Meadow Rush

Lupinus polyphyllus Lupine

Lycopus asper Western Water Horehound

Lycopus europaeus European Water Horehound

Lysimachia nummularia Moneywort

Lysimachia vulgaris Yellow Loosestrife

Lythrum salicaria Purple Loosestrife

Marsilea quadrifolia European Water Clover

Mentha aquatica Watermint

Mentha gracilis Creeping Whorled Mint, Gingermint

Mentha spicata Spearmint

Myosotis scorpiodes True Forget-Me-Not

Myosoton aquaticum Giant Chickweed

Myriophyllum spicatum Eurasian Watermilfoil

Najas marina Spiny Naiad

Najas minor Brittle Waternymph

Nasturtium officianale Water-cress

Nymphoides peltata Yellow Floating Heart

Pluchea odorata succulenta Sweetscent

Pluchea odorata odorata Marsh Fleabane

Poa trivialis Rough-stalked Meadow Grass

Polygonum persicaria Lady’s Thumb, Smartweed, Spotted Knotweed

Potamogeton crispus Curlyleaf Pondweed

Puccinellia distans Weeping Alkali Grass

Rorippa sylvestris Creeping Yellow Cress
16

Rumex longifolius Yard Dock

Rumex obtusifolius Bitter Dock

Salix alba White Willow

Salix fragilis Crack Willow

Salix purpurea Purple Willow

Solanum dulcamara Bittersweet Nightshade

Solidago sempervirens Seaside Goldenrod

Sparganium glomeratum Bur Reed

Trapa natans Water Chestnut

Typha angustifolia Narrow-Leaved Cattail

Veronica beccabunga European Brooklime

Platyhelminthes

Bothriocephalus acheilognathi Asian Tapeworm

Dactylogyrus amphibothrium Monogenetic Fluke

Dactylogyrus hemiamphibothrium Monogenetic Fluke

Dugesia polychroa Flatworm

Ichthyocotylurus pileatus Digenean Fluke

Neascus brevicaudatus Digenean Fluke, Trematode

Scolex pleuronectis Cestode

Timoniella sp. Digenean Fluke, Trematode

Protozoans

Acineta nitocrae Suctorian Ciliate

Glugea hertwigi Microsporidean Parasite

Heterosporis sp. Microsporidean Parasite

Myxobolus cerebralis Salmonid Whirling Disease

Psammonobiotus communis Testate Amoeba

Psammonobiotus dziwnowi Testate Amoeba

Psammonobiotus linearis Testate Amoeba

Sphaeromyxa sevastopoli Myxosporean Parasite

Trypanosoma acerinae Flagellate Parasite

Viruses

Novirhabdovirus p. genotype IV 

sublineage b

Viral Hemorrhagic Septicemia Virus (VHS)

Ranavirus Largemouth Bass Virus (LMBV)

Rhabdovirus carpio Spring Viremia of Carp (SVC)

3. Nonindigenous Species Management

There are two fundamentally different regulatory approaches to address nonindigenous species. 

One approach is to address a particular listed or known species, and another approach targets

certain pathways by which a variety of species may be introduced (Corn and Johnson 2013). 

Both regulatory approaches are employed in the Great Lakes region at both the federal and state 

levels.
17

3.1 Regulations Targeting Pathways

A variety of state and federal regulations target the intentional and unintentional movement of 

species via various pathways without mentioning any particular species by name. 

Ballast water regulations are aimed primarily at interdicting species which may be transported 

and introduced via the shipping ballast pathway. Presently, any ballast-related water in 

essentially all vessels over 79 feet in length that enter the Great Lakes through the St. Lawrence 

Seaway must have a salinity over 30 ppt, obtained either by mid-ocean exchange or exchange 

conducted at approved alternate locations. Any ballast tanks containing water not meeting that 

criteria are sealed and checked again as the ship leaves the Great Lakes. The United States Coast 

Guard (USCG) and the United States Environmental Proection Agency (USEPA) are 

implementing regulations under both the National Invasive Species Act (1996) and the Clean 

Water Act that will require treatment of ballast water prior to discharge to any United States

waters. Discharged ballast water will have to meet numeric discharge standards for live organism 

content (DHS 2012). Treatment systems must be approved by the USCG for use in United States

waters and several treatment systems have been approved on a short-term (five year) basis under 

the USCG "Alternative Management System" Program.

3.2 Federal Regulations Targeting Specific Species

A handful of aquatic invasive species established in the Great Lakes are called out by name for 

stringent regulation under federal laws. These species include Zebra and Quagga Mussels

(Dreissena spp.), Sea Lamprey (Petromyzon marinus), noxious weeds (Sparganium spp.), and 

viruses (Viral Hemorrhagic Septicemia (VHS)).

The injurious wildlife provision of the Lacey Act (1900) bans import and shipment of listed 

species between the continental United States, the District of Columbia, Hawaii, the 

Commonwealth of Puerto Rico, or any possession of the United States. A 1990 amendment to 

the Lacey Act prohibits the possession and transportation of Zebra Mussel (Dreissena 

polymorpha) and Quagga Mussels (D. rostriformis bugensis) in the United States unless intended 

for scientific purposes (by permission of the Secretary of the Interior).

The Great Lakes Fishery Commission was formed in 1955 in part to control Sea Lamprey. The 

commission, in cooperation with Fisheries and Oceans Canada, the United States Fish and 

Wildlife Service, and the United States Army Corps of Engineers participates in Sea Lamprey 

control on the Great Lakes. Fisheries management on the Great Lakes, including management of 

nonindigenous fishes (as well as any factors affecting fish stocks of common concern), is 

coordinated through the Great Lakes Fishery Commission under the Convention on Great Lakes 

Fisheries of 1954, though the states have ultimate authority for management within their borders.
18

The Federal Noxious Weed Act (1974) authorizes management of federally listed noxious 

weeds. Under the synonym, Sparganium erectum, S. glomeratum is federally listed as a noxious 

weed (Code of Federal Regulations, Title 7, 360.200).

The Animal Damage Control Act (1931) authorizes the United States Department of Agriculture 

Animal and Plant Health Inspection Service (APHIS) to control damage caused by wildlife to 

agricultural interests (including aquaculture), and the Federal Seed Act (1939) adds regulation of 

noxious weed seeds. Viral Hemmorhagic Septicemia (VHS) is an internationally-reportable fish 

disease. Permits from APHIS and health inspections were required for interstate movement of 

VHS-susceptible fish from VHS-infected or at risk states to non-infected states during 2006-

2014. Post-2014, the federal order was removed as duplicative of state regulations. A United 

States Department of Agriculture import permit and a veterinary health certificate must 

accompany imports of live fish, fertilized eggs, and gametes of species susceptible to Spring 

Viremia of Carp including Common Carp (Cyprinus carpio), Koi (C. carpio koi), Grass Carp 

(Ctenopharyngodon idella), Silver Carp (Hypophthalamicthys molitrix), Bighead Carp (H. 

nobilis), Crucian Carp (Carassius carassius), Goldfish (Carassius auratus), Tench (Tinca tinca), 

Ide (Leuciscus idus), and Wels Catfish (Silurus glanis) (USDA APHIS 2012).

3.3 State Regulations Targeting Specific Species

A much broader array of established nonindigenous aquatic species are targeted for regulation in 

state laws in the Great Lakes region. Most states in the Great Lakes take a blacklist approach to 

regulation of aquatic invasive species in which species are regulated only if named in specific 

legislation/regulation. Illinois takes a whitelist approach to regulation of aquatic invasive species 

in which all aquatic species – including fish, reptiles, amphibians, mollusks, crustaceans, algae, 

aquatic plants, and aquatic invertebrates – are regulated unless approved. The whitelist approach 

results in the effective prohibition on aquaculture, transport, stocking, import and possession of a 

large number of species that are not specifically named in legislation. Illinois additionally 

blacklists some species as ‘injurious’ with more stringent regulation and penalties.

3.3.1 Fish pathogens

Five species of fish pathogens, including two bacteria (Renibacterium salmoninarum responsible 

for Bacterial Kidney Disease and Aeromonas salmonicida responsible for furunculosis), a 

myxosporean (Myxobolus cerebralis responsible for Whirling Disease), and two viruses 

(Novirhabdovirus sp. responsible for Viral Hemorrhagic Septicemia (VHS) and Ranavirus sp.

responsible for Large Mouth Bass Virus) are explicitly targeted for regulation by the Great Lakes 

states. Four of these species (all but A. salmonicida) are considered to have high environmental 

impact to the Great Lakes. A. salmonicida is targeted primarily because it is considered a threat 

to hatchery operations for rearing salmonids. Regulations focus on transportation, stocking, and 

baitfish as the primary vectors for the movement of fish pathogens.
19

Table 1. Great Lakes States that regulate fish pathogens.

MN WI IL IN MI OH PA NY

Aeromonas 

salmonicida

√ √ √ √ √ √ √ √

Renibacterium 

salmoninarum

√ √ √ √ √ √ √ √

Myxobolus 

cerebralis

√ √ √ √ √ √ √ √

Novirhabdovirus 

sp. genotype IV 

sublineage b

√ √ √ √ √ √ √ √

Ranavirus sp. √ √ √ √

Regulations that target Novirhabdovirus sp. (VHS) focus on transport. Transportation of VHSsusceptible species is prohibited from New York, Pennsylvania, Ohio, Michigan, Indiana, Illinois

(including movement of natural water from infested waters), Wisconsin, Minnesota, Ontario, and

Quebec unless certain conditions are met (see below; USDA APHIS 2008). Movement of VHSsusceptible species is permitted from the two infected Canadian provinces to the United States if 

the shipment meets certain requirements and is imported under an APHIS permit for direct 

slaughter, or during catch and release fishing activities (USDA APHIS 2008). Movement of 

VHS-susceptible species is permitted between VHS-infected or at-risk states as long as fish are 

sent directly to state-inspected slaughter facilities that discharge waste water to a municipal

sewage system that includes disinfection, or discharge to a non-discharging pond or a settling 

pond that disinfects according to all applicable United States Environmental Protection Agency 

(USEPA) and state regulatory criteria, and are accompanied by a valid VS 1-27 (Permit for 

Movement of Restricted Animals) form issued by an APHIS area office. Remains from slaughter 

facilities must be rendered or composted (USDA APHIS 2008). Interstate movement of VHSsusceptible fish is permitted from VHS-infected or at risk states to non-infected states as long as 

the fish are accompanied by appropriate state, tribal, or federal documentation stating the fish 

have tested negative for the virus (USDA APHIS 2008). Movement of VHS-susceptible species 

may also be permitted to state, federal, or tribal-authorized research and diagnostic facilities 

which meet containment and discharge requirements provided that the fish are accompanied by a 

valid VS 1-27 form issued by an APHIS area office and the remains are disposed of as medical 

waste adhering to all applicable USEPA and state regulatory criteria (USDA APHIS 2008).

Regulations targeting Renibacterium salmoninarum, Myxobolus cerebralis, Aeromonas 

salmonicida, and Ranavirus sp. focus primarily on stocking. The Great Lakes Fish Disease 

Control Policy and Model Program has prohibited stocking the Great Lakes and their tributaries 

with fish from whirling disease-infected farms. Fish imported into north-central-region states 

must be certified free of whirling disease in order to obtain import permits (Faisal and Garling 

2004). Ohio requires out-of-state source facilities to document annual salmonid fish, egg, and 
20

sperm health inspections for one year prior to importation (NCRAC 2010ab). Ohio further 

requires source facilities outside the Great Lakes basin to document annual health inspections 

showing no furunculosis, Bacterial Kidney Disease (BKD), or whirling disease occurrences for 

the previous five years prior to importing salmonids to the Lake Erie watershed (Baird 2005). 

Indiana requires source facilities within the Great Lakes basin to document they have been free 

of whirling disease for three consecutive years prior to importing salmonid stock. Source 

facilities outside the basin must document that salmonid stocks have been free of whirling 

disease, BKD, and furunculosis consecutively since 2002 (Baird 2005). Salmonids found 

carrying the pathogen, but asymptomatic, can be sold within states if source facilities are within 

the Great Lakes basin (Baird 2005). Michigan requires imported aquacultured fish to be 

accompanied by either an official interstate health certificate, official interstate certificate of 

veterinary inspection, or a fish disease inspection report. Importing of aquacultured fish is 

prohibited from source facilities with a record of an emergency disease within the past two years. 

Fish must be certified free of Largemouth Bass Virus (LMBV) if they are imported from nonMichigan source facilities and intended for stocking in public waters (NCRAC 2010ab). 

Michigan requires source facilities to document salmonid stocks have been whirling disease free 

for two consecutive years prior to importation, while Wisconsin requires one. Wisconsin requires 

source facilities to document fish health inspections targeting LMBV prior to importing live fish 

and eggs (NCRAC 2010ab). Both Michigan and Wisconsin also target BKD. Illinois requires 

source facilities of any species of live fish, eggs, and sperm to document they are disease free 

prior to importation (NCRAC 2010ab) and the testing of these facilities must be to the World 

Organization for Animal Health (OIE) or the American Fisheries Society (AFS) Bluebook 

standards and have appropriate HACCP in place. Further, Illinois restricts the import of live fish 

and gametes from certain geographic locations. Illinois and Minnesota also require imported 

salmonid health inspections. Minnesota allows the importation of whirling disease-infected eggs, 

if prior egg treatments are approved (Baird 2005). Prior to placing fish in New York waters, a 

fish health certification report must document that the fish are furunculosis, BKD, and VHS free.

All eight Great Lakes states have instated similar baitfish regulations to control the spread of 

Myxobolus cerebralis and Novirhabdovirus sp. New York, Pennsylvania, Michigan, Illinois, 

Wisconsin, and Minnesota have instated similar baitfish regulations to control the spread of 

furunculosis, BKD, and other fish pathogens. New York regulations include that bait harvested 

from inland waters for personal use is only permitted to be used within the same body of water 

from which it was taken and cannot be transported overland (with the exception of Osmerus 

mordax, suckers (Catastomidae), Alosa pseudoharengus, and Alosa aestivalis). Once transported, 

baitfish cannot be replaced to their original body of water (NYSDEC 2012a). Live or frozen bait 

harvested from inland New York waters for commercial purposes is only permitted to be sold or 

possessed on the same body of water from which it was taken and cannot be transported over 

land unless under a permit and/or accompanied by a fish health certification report. Bait that is 

preserved and packaged by any method other than freezing, such as salting, can be sold and used 

wherever the use of bait fish is legal as long as the package is labeled with the name of the 
21

packager-processor, the name of the fish species, the quantity of fish packaged, and the means of 

preservation (NYSDEC 2012a). Certified bait may be sold for retail and transported overland as 

long as the consumer maintains a copy of a sales receipt that contains the name of the selling 

vendor, date sold, species of fish sold, and quantity of fish sold. Bait that has not been certified 

may still be sold, but the consumer must maintain a sales receipt containing the body of water 

where the bait fish was collected and a warning that the bait cannot be transported by motor 

vehicle. Bait sold for resale requires a fish health certification along with a receipt that contains 

the name of the selling vendor, date sold, species of fish sold, and quantity of fish sold, which 

must be kept for 30 days or until all bait is sold (NYSDEC 2012a).

3.3.2 Fish

Pacific salmon (Oncorhynchus gorbuscha, O. kisutch, O. mykiss, O. nerka, O. tshawytscha) were 

introduced to the Great Lakes in the 1960s to manage Alewife (Alosa pseudoharengus)

populations and support a fishery (Tody and Tanner 1966). Soon after, the multi-million dollar 

Great Lakes Pacific salmon sportfishery was established and is now supports one of the largest 

economic sectors in the region. Therefore, Pacific salmon management objectives are designed

to maintain or enhance the health and stability of the fisheries. Pacific salmon management is

extremely diverse, integrated, and cascading and is therefore these are the most heavily regulated 

species (direct and indirectly) in the Great Lakes. Typically, Pacific salmon regulations are not 

species specific, but rather regulate the salmonid fisheries as a whole. Salmo trutta, Osmerus 

mordax, Morone americana, Lepomis microlophus, and Cyprinus carpio are other nonnative 

species which are managed at least in part to support fisheries (moderate to high beneficial 

species). Alosa pseudoharengus is the prime prey of Pacific salmonids and is managed in part to 

support the Pacific salmonid fisheries. 

Table 2. Nonindigenous fishes managed primarily as beneficial species in Great Lakes States.

MN WI IL IN MI OH PA NY

Lepomis microlophus √ √

Notropis buchanani √

Oncorhynchus gorbuscha √ √ √ √ √ √ √ √

Oncorhynchus kisutch √ √ √ √ √ √ √ √

Oncorhynchus mykiss √ √ √ √

Oncorhynchus tshawytscha √ √ √ √ √ √ √ √

Salmo trutta √

Great Lakes states and provinces have their own specific fishing regulations. Generally, the 

overall goals and objectives of Pacific salmon (and other) fishing regulations are the same 

throughout the region. Fishing regulations include daily and season bag limits, size limits, 

permitted baitfish, manner of take (i.e., snagging or hook and line), and designated season dates 
22

(See New York State DEC, Pennsylvania F&BC, Ohio DNR, Michigan DNR, Indiana DNR,

Illinois DNR, Minnesota DNR, Wisconsin DNR websites for specific fishing regulations). Note 

that Ontario takes a whitelist approach to baitfish – all unlisted species are prohibited to use as 

bait.

The Ghost Shiner (Notropis buchanani) is native south of the Great Lakes basin and is listed as 

an endangered species in Pennsylvania. Pennsylvania state law prohibits the catch, take, kill, 

possession, and import to or export from Pennsylvania, sale or offer of sale or purchase of any 

individual of an endangered species, alive or dead, or any part thereof, without a special permit, 

(58 PA Code § 75.1). In locations where it is invasive to the Great Lakes basin, the 

environmental impacts of N. buchanani are unknown and socioeconomic impacts are low. No 

other state in the Great Lakes region regulates N. buchanani.

In addition to the above regulations on fish transport, stocking, baitfish (which target fish 

pathogens), fishing regulations (targeted at sustaining fisheries), and endangered species 

protections, there are state regulations for 14 species of nonindigenous fish which focus on 

controlling or managing fish as invasives – plus one species that is effectively restricted by a 

whitelist approach. Four of these species, Alewife (Alosa pseudoharengus), Common Carp

(Cyprinus carpio), Rainbow Smelt (Osmerus mordax), and White Perch (Morone americana)

each have favorable benefits as well as negative environmental impacts. The remaining 10

species include a mix of species with high to moderate impacts as well as a few for which impact 

has not been adequately assessed. 
23

Table 3. Nonindigenous fishes regulated primarily as invasives in Great Lakes states.

MN WI IL IN MI OH PA NY

Alosa aestivalis Restricted Prohibited

exotic

Restricted

Alosa 

pseudoharengus

Regulated Approved Prohibited

exotic

Restricted

Apeltes 

quadracus

Restricted

Carassius 

auratus

Regulated Restricted

invasive

Approved Prohibited

bait

Restricted

Cyprinus carpio Regulated Restricted

invasive

Approved Regulated Prohibited

exotic

Prohibited

bait

Restricted

Enneacanthus 

gloriosus

Restricted

Gambusia affinis Prohibited Native

Gymnocephalus 

cernua

Prohibited Restricted

invasive

Injurious Regulate Prohibited Prohibited Prohibited

Lepisosteus 

platostomus

Native Permit

Required

Misgurnus 

anguillicaudatus

Restricted

Aquarium

Restricted Prohibited

Morone 

americana

Prohibited Restricted Regulated Prohibited

Neogobius 

melanostomus

Prohibited Injurious Regulated Prohibited Prohibited Prohibited Restricted

Oncorhynchus 

nerka

Restricted

Osmerus mordax Regulated Restricted

invasive

Injurious Regulated

sportfish

Commercial

unrestricted

Regulated

commercial

Restricted

Petromyzon 

marinus

Prohibited Prohibited Restricted

Proterorhinus 

semilunaris

Prohibited Restricted

invasive

Injurious Regulated Prohibited Prohibited Prohibited

Scardinius 

erythrophthalmus

Prohibited Prohibited Injurious Regulated Prohibited Prohibited Prohibited

Gambusia affinis and Scardinius erythophthalmus are prohibited in Wisconsin, meaning that no 

person may transport, possess, transfer, or introduce the species without authorization (WI
24

Administrative Code § NR 40.04). Gymnocephalus cernua, Proterorhinus semilunaris, Osmerus 

mordax, Cyprinus carpio and Carassius auratus are restricted invasive species, and therefore 

cannot be transported, possessed, transferred, or introduced without a permit (WI Administrative

Code § NR 40.05). Oncorhynchus mykiss, Salmo trutta, and Lepomis microlophus are listed as 

restricted species under the definition “nonnative fish in the aquaculture industry” (WI

Administrative Code § NR 40.02, 40.05) and therefore cannot be transported, possessed, 

transferred, or introduced without a permit. Misgurnus anguillicaudatus is restricted as a 

nonnative viable fish species in the aquarium trade (WI Administrative Code § NR 40.02, 40.05). 

In Minnesota, Petromyzon marinus, Gymnocephalus cernua, Morone americana, Scardinius 

erythrophthalmus, Proterorhinus semilunaris, and Neogobius melanostomus are prohibited 

invasive species, meaning it is unlawful (a misdemeanor) to possess, import, purchase, transport, 

or introduce an organism except under permit for control, research, or education (MN

Administrative Rules § 6216.0250). Alosa pseudoharengus, Cyprinus carpio, Osmerus mordax,

and Carassius auratus are regulated invasive species (MN Administrative Rules § 6216.0260 

Regulated), making it illegal to introduce the species without a permit (MN Administrative Rules 

§ 6216.2060, MN Administrative Rules § 6216.0265). 

In Ohio, it is unlawful for any person to possess, import, or sell live Petromyzon marinus, 

Neogobius melanostomus, Proterorhinus semilunaris, Gymnocephalus cernua, or Morone 

americana (Ohio Administrative Code §1501:31-19). A class B aquaculture permit is required to 

engage in propagation, culture, or sale of Shortnose Gar (Lepisosteus platostomus), and two 

levels of escapement prevention are required if cultured in the Lake Erie drainage basin (Ohio 

Administrative Code § 1501:31-39-01). In Ohio it is illegal for any person to possess, import or 

sell exotic species of fish including Alosa aestivalis, Alosa pseudoharengus, and Cyprinus carpio

and/or hybrids for introduction or to release into any body of water that is hydologically 

connected to public waters, or waters of the state, without first having obtained permission (Ohio 

Administrative Code §1501:31-19). In Ohio, Osmerus mordax is defined as a commercial fish 

and an unrestricted species under Ohio Administrative Code 1501 § 31-1-02. Commercial fish 

are permitted to be taken, possessed, bought, or sold unless otherwise restricted in Ohio code. 

In Indiana, Gymnocephalus cernua, Neogobius melanostomus, Proterorhinus semilunaris, and 

Morone americana are classified as exotic fish (312 IN Administrative Code § 9-6-7), meaning 

except as otherwise provided, no individual can import, possess, propagate, buy, sell, barter, 

trade, transfer, loan, or release into public or private waters live fish, recently hatched juveniles, 

viable eggs, or genetic material. Indiana (312 IN Administrative Code § 9-6-8) prohibits the use 

of carp as bait. Indiana also has no bag limit for Cyprinus carpio and has legalized spearfishing, 

bowfishing and snaring for this species. In Indiana, Osmerus mordax sport fishing season on 

Lake Michigan is defined as March 1-May 30, with capture allowed only by the use of dip nets, 

seines, or nets (312 IN Administrative Code § 9-7-2); otherwise, there is no bag limit, possession 

limit, or size limit as defined under 312 IN Administrative Code § 9-7-14. 
25

In Pennsylvania (58 PA Code §71.6), it is illegal to possess, import, or introduce Neogobius 

melanostomus, Proterorhinus semilunaris, Scardinius erythophthalmus or Gymnocephalus 

cernua. Further, it is unlawful to sell, purchase, offer for sale, or barter for live Ruffe (58 PA 

Code § 63.46). Ruffe may not be transported from another state, province, or country into 

Pennsylvania, liberated into a Pennsylvania watershed, or transferred between Pennsylvania 

waters without written permission from the Pennsylvania Fish and Boat Commission under 58 

PA Code § 73.1, Pennsylvania (58 PA Code §63.44) prohibits the use of carp and Goldfish

(Carassius auratus) as bait. The use of commercial trap nets under license to capture Rainbow 

Smelt (Osmerus mordax) is regulated by Pennsylvania Administrative Code § 69.33. 

In Michigan, Misgurnus anguillicaudatus, Gymnocepalus cernua, Scardinius erythophthalmus, 

Proterorhinus semilunaris, and Neogobius melanostomus are prohibited species (MI NREPA 

451 § 324.41301). Both introduction and possession of these species is prohibited except for 

education, research, or identification purposes (MI NREPA 451 § 324.41303). A violation 

involving a prohibited species is a felony, and a knowing introduction with intent to harm is 

punishable with up to five years imprisonment and a $2,000 to $1,000,000 fine (MI NREPA § 

324.41309). Lepomis microlophus and Oncorhynchus mykiss are listed as approved species for 

aquaculture production in Michigan (Aquaculture Development Act; MCL § 286.875).

Illinois takes a whitelist approach to invasive fish species. Any non-native species not explicitly 

listed as approved for aquaculture, transportation, stocking, importation, and/or possession is 

automatically considered to be restricted. Alosa pseudoharengus, Carassius auratus, and 

Cyprinus carpio are approved. While not native to Lake Michigan, Gambusia affinis, 

Lepisosteus platostomus, Esox niger, Ictiobus cyprinellus, and Phenacobius mirabilis are native 

elsewhere in Illinois and so are not restricted. Though not called out by name, Alosa aestivalis, 

Misgurnus anguillicaudatus, Morone americana, Oncorhynchus nerka, Apeltes quadracus, and

Enneacanthus gloriosus are considered restricted. Gymnocephalus cernua, Proterorhinus 

semilunaris, Neogobius melanostomus, and Scardinius erythophthalmus are listed as injurious 

species under Illinois Administrative Code 17 § 805.20. It is unlawful to possess, propagate, buy, 

sell, barter, or offer to be bought, sold, bartered, transported, traded, transferred, or loaned an 

injurious species to any person or institution unless a permit is obtained from the Illinois DNR 

(IL Administrative Code 17 § 805.30). Rainbow Smelt (Osmerus mordax) is regulated as a 

sportfish; the season for O. mordax is defined as March 1-April 30 under Illinois Administrative

Code 17-1 § 810.10. 

New York, under federal law, is required to follow the Interstate Fishery Management Plan for 

Shad and River Herring established by the Atlantic States Marine Fisheries Commission 

(Enacted May 2009; ASMFC). The New York Department of Environmental Conservation is

therefore forced to close any non-sustainable commercial and recreational fisheries by January 1, 

2012 until it can prove New York fisheries are self-sustaining (NYSDEC 2012b). New York 

prohibits or restricts the use of several species of nonindigenous fish as bait including Alosa 

aestivalis, Alosa pseudoharengus, Osmerus mordax, Carassius auratus larvae, Petromyzon 
26

marinus larvae, carp (presumed all species), and Neogobius melanostomus (6 NY Code, Rules, 

and Regulations (NYCRR) § Part 10, Paragraph 10.1(c)(3), 6 NYCRR § Part 19, NY 

Environmental Conservation Law § 11-1315, 6a). Goldfish larvae taken in nets operated 

pursuant to baitfishing are to be destroyed immediately (NY Environmental Conservation Law § 

11-1315). 

3.3.3 Mollusks

Six species of mollusks are state-regulated and an additional 11 species are effectively regulated 

due to Illinois’ whitelist approach. Of these, only Zebra Mussels are regulated in all eight Great 

Lakes states, though Quagga Mussels are federally regulated and thus are effectively regulated in 

all eight states. This list includes a mix of species with high impact (Dreissenid mussels), 

moderate impact (Corbicula fluminea and Potamopyrgus antipodarum), and unknown impact 

species (Cipangopaludina spp.). 

Table 4. Great Lakes states regulating nonindigenous mollusks.

MN WI IL IN MI OH PA NY

Cipangopaludina chinensis 

malleata

√ √ √

Cipangopaludina japonica √ √

Corbicula fluminea √ √ √

Dreissena polymorpha √ √ √ √ √ √ √ √

Dreissena rostriformis 

bugensis

√ √ √ √ √ √

Elimia virginica √

Gillia altilis √

Lasmigona subviridis √

Pisidium amnicum √

Pisidium henslowanum √

Pisidium moitessieranum √

Pisidium supinum √

Potamopyrgus antipodarum √ √ √

Radix auricularia √

Sphaerium corneum √

Valvata piscinalis √

Corbicula fluminea and Potamopyrgus antipodarum are prohibited species in Wisconsin (WI 

Administrative Code § NR 40.04). Cipangopaludina chinensis is a restricted species in 

Wisconsin (WI Administrative Code § NR 40.05). In addition to the federal restrictions, in 
27

Wisconsin, it is unlawful to transport, transfer, or introduce Zebra Mussels (Dreissena 

polymorpha) (WI Administrative Code § NR 40.05). 

Potamopyrgus antipodarum are listed as a prohibited species in Minnesota (MN Administrative 

Rules § 6216.0250). Cipangopaludina spp. are regulated invasive species in Minnesota (MN

Administrative Rules § 6216.0260). In addition to federal regulations, in Minnesota it is unlawful 

to place or attempt to place into state water a watercraft, trailer, or plant harvesting equipment 

that has attached Zebra Mussels (MN Statuses § 84D.10). Persons leaving the state are required 

to drain boats and related equipment during transportation on a public road (MN Statuses § 

84D.10, MN Administrative Rules 6216.0500, Kaminski Leduc 2011).

In Indiana, an individual may not import, possess or release Asiatic clams (including Corbicula 

fluminea) or dreissenid mussels (Dreissena polymorpha and Dreissena rostriformis bugensis)

into public or private waters (312 IN Administrative Code § 9-9-3).

Illinois also applies the whitelist approach to non-native mollusks. Thus Corbicula fluminea, 

Cipangopaludina sp., Potamopyrgus antipodarum, Sphaerium corneum, and Valvata piscinalis

are restricted, even though they are not explicitly mentioned in legislation. Lasmigona subviridis,

Pisidium amnicum, P. henslowanum, P. moitessierianum, P supinum, Elimia virginica, Radix 

auricularia, and Gillia altilis are also not explicitly mentioned in Illinois’ legislation, but are not 

currently found in Illinois waters – under the whitelist approach import of these species to 

Illinois is prohibited as is possession in the state. Viviparus georgianus and Bithynia tentaculata

are approved species in Illinois. Illinois lists Zebra Mussels as injurious species (IL

Administrative Code 17 § 805.20). 

In addition to federal regulations, the following states have additional restrictions for mollusks 

specific to Dreissena spp. In Pennsylvania, it is unlawful to possess, introduce, import, transport, 

sell, purchase, offer for sale, or barter Zebra Mussels (58 PA Code § 63.46, 58 PA Code § 71.6, 

58 PA Code § 73.1). In Michigan, Zebra Mussels are a restricted species (MI Compiled Laws § 

324.41301) and therefore cannot be possessed unless it is to identify, eradicate, or control the 

species (MI Compiled Laws § 324.41303). In Ohio, it is unlawful to possess, import, or sell 

Zebra Mussels (Ohio Administrative Code § 1501:31-19-01). In New York, it is unlawful to 

intentionally release Zebra Mussels into state waters (NY Environmental Conservation Law § 11-

0507). 

3.3.4 Waterfleas and Mysids

Three species of established nonindigenous waterfleas as well as one mysid are regulated by 

name by select Great Lakes states. These include two high impact raptorial waterfleas 

(Bythotrephes longimanus and Cercopagis pengoi) and two relatively recent invaders whose 

impact has not been assessed (Daphnia lumholtzi and Hemimysis anomala). An additional 13

species are covered solely by Illinois’ whitelist approach.
28

In Wisconsin, Bythotrephes longimanus, Cercopagis pengoi, Daphnia lumholtzi, and Hemimysis 

anomala are prohibited invasive species. With certain exceptions, it is unlawful to transport, 

possess, transfer, or introduce a prohibited invasive species in Wisconsin (WI Administrative

Code § NR 40.04). In Minnesota, Bythotrephes longimanus is a regulated invasive species (MN

Administrative Rules § 6216.0260); it is legal to possess, sell, buy, and transport regulated 

invasive species, but no person may introduce a regulated invasive species without a permit (MN

Administrative Rules § 6216.0265 Subpart 1). 

Illinois’ whitelist approach also applies to crustaceans, therefore Bythotrephes longimanus, 

Cercopagis pengoi, Daphnia lumholtzi, Hemimysis anomala, Eurytemora affinis, Heteropsyllus 

nr. nunni, Nitokra hibernica, Schizopera borutzkyi, Bosmina maritima, Echinogammarus 

ischnus, and Gammarus tigrinus which are not approved, are considered restricted. Cyclops 

strenuous, Megacyclops viridis, Nitokra incerta, Skistodiaptomus pallidus, and Daphnia galeata 

galeata are also not approved, but not currently found in Illinois waters – possession and 

importation of these species is prohibited. 
29

Table 5. Great Lakes states regulating nonindigenous crustacean species.

MN WI IL IN MI OH PA NY

Bosmina coregoni √

Bythotrephes 

longimanus

√ √ √

Cercopagis 

pengoi

√ √

Daphnia galeata 

galeata

√

Daphnia 

lumholtzi

√ √

Eubosmina 

maritima

√

Cycops strenuuus √

Eurytemora 

affinis

√

Heteropsyllus nr.

Nunni

√

Megacyclops 

viridis

√

Nitokra hibernica √

Nitokra incerta √

Schizopera 

borutzkyi

√

Skistodiaptomus 

pallidus

√

Echinogammarus 

ischnus

√

Gammarus 

tigrinus

√

Hemimysis 

anomala

√ √

3.3.5 Plants

In addition to the federal regulations applicable to the noxious weed Sparganium glomeratum, 

the Great Lakes states regulate 26 species of nonindigenous aquatic plants which are established 

in the basin. Seventy three percent of the listed plants have documented moderate to high 

environmental and or socioeconomic impacts. An overlapping 35% of plants have documented 

benefits. No individual plant species is regulated in all eight Great Lakes states. Some species 

(e.g., Black Alder, Alnus glutinosa), which are restricted in one state, are recommended in 

another. In Pennsylvania, Sweetscent (Pluchea odorata) (native to the eastern United States) is 

listed as an endangered species; no other Great Lakes state regulates this species.
30

Table 6. Great Lakes states regulating nonindigenous plant species.

MN WI IL IN MI OH PA NY

Alnus glutinosa √ √ Approved

Alopecurus geniculatus Restricted

Butomus umbellatus √ √ Injurious √ √ √ √ √

Cabomba caroliniana √ √ Restricted √ √

Carex acutiformis Restricted

Carex disticha Restricted

Chenopodium glaucum ?

Cirsium palustre √ √ Restricted √ √

Conium maculatum √ Restricted √

Echinochloa crus-galli √ ?

Epilobium hirsutum √ ? √ √

Frangula alnus √ √ Regulated

Exotic Weed

√

Glyceria maxima √ Restricted √

Hydrocharis morsus-ranae √ √ Injurious √ √ √

Iris pseudacorus √ Injurious √ √ √ √

Juncus compressus Restricted

Juncus gerardii Restricted

Juncus inflexus Restricted

Lycopus europaeus Restricted

Lysimachia nummularia ? √

Lysimachia vulgaris ? √

Lythrum salicaria √ √ Regulated

Exotic Weed

√ √ √ √

Marsilea quadrifolia Restricted

Myosotis scorpiodes * ?

Myriophyllum spicatum √ √ Injurious √ √

Najas marina √ Approved

Najas minor √ √ Injurious √ √

Nasturtium officianale √ Approved

Nymphoides peltata √ Injurious √ √ √

Pluchea odorata Restricted*** Native

∗∗

Potamogeton crispus √ √ Injurious √ √ √

Rorippa sylvestris Restricted

Solanum dulcamara ? √

Solidago sempervirens Restricted

Sparganium glomeratum √ Injurious √

Trapa natans √ √ Injurious √ √ √
31

Typha angustifolia √ Approved √

Veronica beccabunga Restricted

*Proposal pending in WI to list Myosotis scorpiodes as a restricted species.

** In PA, Pluchea odorata is listed as an endangered species.

***Pluchea odorata succulenta is native to parts of Illinois and not regulated. Pluchea odorata 

odorata is not native and restricted. 

? Plants which are not obligate aquatic species (e.g., facultative and wetland plants) may not be 

considered ‘aquatic’ and so may not be covered under the IL Whitelist approach.

Cirsium palustre, Cabomba caroliniana, Epilobium hirsutum, Glyceria maxima, Hydrocharis 

morsus-ranae, Myriophyllum spicatum (including hybrids and variants), Najas minor, 

Nymphoides peltata, and Trapa natans are prohibited species in Wisconsin. Wisconsin law 

exempts some species in some counties (generally where already very well established) from the 

full restrictions otherwise associated with prohibited species. Lythrum salicaria is designated 

both as a restricted species (WI Administrative Code § NR 40.05) and as an invasive aquatic 

plant (WI Administrative Code § NR 109.07). Butomus umbellatus, Frangula alnus, 

Potamogeton crispus, and Typha angustifolia are listed as restricted species, and may not be 

transported, transferred, or introduced into any ecosystem. The Robert W. Freckman Herbarium 

of the University of Wisconsin lists an eradication notice for Poison Hemlock (Conium

maculatum). However, as of 2007 it was not illegal to sell C. maculatum in Wisconsin (Annen 

2007). However, the possession, transportation, transfer, or introduction of C. maculatum is 

restricted in most Wisconsin counties (WIDNR 2010a). Even though it is not restricted or 

prohibited, the Wisconsin Department of Natural Resources acknowledges Nasturtium officinale

as being highly invasive and recommends its eradication upon detection (Robert W. Freckman 

Herbarium 2012). Alnus glutinosa is listed as a “plant to avoid” in Wisconsin’s planting guide. 

Butomus umbellatus, Hydrocharis morsus-ranae, Lythrum salicaria (including all cultivars) 

Myriophyllum spicatum (including all hybrids and variants), Najas minor, Potamogeton crispus, 

Sparganium glomeratum, and Trapa natans are listed as prohibited species in Minnesota (MN 

Administrative Rules § 6216.0250). Frangula alnus is listed as a restricted noxious weed, and 

the importation, sale, or transport this plant is illegal (MNDNR 2009). Iris pseudacorus is 

established and considered a “moderate threat” to local ecosystems in Minnesota; this has led it 

to be classified as a restricted species that cannot be planted/released without a permit (GLPANS 

2008, Minnesota Invasive Species Advisory Council 2009). Cabomba caroliniana can be 

possessed, sold, bought, and transported, but it is illegal to release it into the environment (MN 

DNR 2013b). Even though it is not present in Minnesota, Cirsium palustre is characterized as a 

severe threat to native ecosystems based on its impact in other locations (Minnesota Invasive 

Species Advisory Council 2009). Echinochloa crus-galli is considered to pose a “minimal” 

threat to ecosystems: it doesn’t pose significant competition with native species, it may 

naturalize, it does not significantly alter ecosystems, and has little possibility of spread within or 
32

to other sites (Minnesota Invasive Species Advisory Council 2009). Najas marina is listed as a 

“species of special concern”; meaning it is extremely uncommon and deserves careful 

monitoring of its status (MN DNR 2013b). Alnus glutinosa is a recommended tree for urban 

environments in Minnesota (Johnson and Himanga 2009).

Trapa natans is prohibited in New York (GLPANS 2008). The New York Invasive Species 

Council assessed Cabomba caroliniana, Frangula alnus, Glyceria maxima, Hydrocharis morsusranae, Iris pseudacorus, Lysimachia nummularia, Lysimachia vulgaris, Nymphoides peltata, and

Potamogeton crispus as having a high risk of causing ecological harm and recommend that their 

use be prohibited (NYISC 2010). The New York Invasive Species Council ranks Najas minor a 

moderate ecological risk and recommends that the species be regulated (NYISC 2010). Cirsium 

palustre and Epilobium hirsutum are also considered to be a medium to high threat species in 

New York (Higman and Campbell 2009, NYISC 2010). Butomus umbellatus is on New York 

State’s Interim Invasive Species Plant List (NYSDEC 2011). 

Butomus umbellatus, Hydrocharis morsus-ranae, Iris pseudacorus, Myriophyllum spicatum,

Najas minor, Nymphoides peltata, Potamogeton crispus, Sparganium glomeratum, and Trapa 

natans are listed as injurious species in Illinois. Lythrum salicaria and Frangula alnus are listed 

as exotic weeds in Illinois (IL Compiled Statuses 525; 10/3 and 10/4) making it illegal to buy, 

sell, or distribute plants, its seeds, or any part without a permit. Illinois’ whitelist approach 

applies to aquatic plants, but the definition of aquatic plant used does not correspond completely 

to the definition used by GLANSIS. Nonindigenous submerged, floating and emergent aquatic 

plants including Alopecurus geniculatus, Cabomba caroliniana, Carex acutiformis, Carex

disticha, Cirsium palustre, Conium maculatum, Juncus compressus, Juncus gerardii, Juncus 

inflexus, Lycopus europaeus, Marsilea quadrifolia, Pluchea odorata odorata, Rorippa sylvestris,

and Veronica beccabunga are clearly restricted under the Illinois whitelist approach. 

Facultative wetland species which can invade aquatic environments but are principally upland 

species may not be restricted – these include Agrostis gigantea, Alnus glutinosa, Chenopodium 

glaucum, Echinochloa crus-galli, Epilobium hirsutum, Glyceria maxima, Impatiens glandulifera, 

Lupinus polyphyllus, Lysimachia nummularia, Lysimachia vulgaris, Mentha aquatica, Mentha 

gracilis,Mentha spicata, Myosotis scorpiodes, Myosoton aquaticum, Poa trivialis, Polygonum 

persicaria, Rumex longifolius, Rumex obtusifolius, Salix alba, Salix fragilis, Salix purpurea, and 

Solanum dulcamara. The New Invaders Watch Program lists Glyceria maxima on its “watch list” 

for Illinois (Maurer 2009). Pluchea odorata succulenta, Lycopus asper, Najas marina, 

Puccinellia distans, Solidago sempervirens, and Typha angustifolia are native to parts of Illinois 

and so not restricted. Najas marina, Typha angustifolia and Nasturtium officianale are approved

for purposes of aquaculture. 

Butomus umbellatus, Cabomba caroliniana, Hydrocharis morsus-ranae, Iris pseudacorus, 

Myriophyllum spicatum, Nymphoides peltata, and Trapa natans are prohibited in Michigan

(GLPANS 2008). Potamogeton crispus and Lythrum salicaria (with an exemption for sterile 
33

cultivars) are restricted (MI NREPA 451 § 324.41301). Cirsium palustre is considered to be a 

medium to high threat species in Michigan (Higman and Campbell 2009). 

In Ohio, Conium maculatum has been designated as a prohibited noxious weed (Ohio Division of 

Natural Areas and Preserves and Nature Conservancy 2000, USDA NRCS 2012). Planting or 

sale of Lythrum salicaria without a permit is (Ohio Revised Code § 927.682), though the director 

may exempt varieties ‘demonstrated not to be a threat to the environment’. Butomus umbellatus, 

Epilobium hirsutum and Iris pseudacorus are classified as“well established invasive plants” by 

the Ohio Department of Natural Resources.

Pennsylvania has designated all nonnative Lythrum species and their cultivars as noxious weeds 

(7 PA Code § 110.1). Butomus umbellatus is labeled as a medium-high threat to native 

ecosystem. In Pennsylvania, Pluchea odorata, which is native to the east coast, is listed as an 

endangered species (USDA NRCS 2012).

Planting, sale, or other distribution of Lythrum salicaria without a permit is also prohibited in 

Indiana (312 IN Code § 14-24-12). Indiana recommends that Black Alder be inter-planted to 

improve soil quality and “protect other valuable trees” (IDNR n.d.) but the state also includes 

Black Alder on a list of invasive exotics plants whose “use in landscaping and re-vegetation 

projects should be avoided or limited when possible” (Homoya 2010). 

3.3.6 Miscellaneous taxa covered only under the whitelist approach – Algae and Aquatic 

Invertebrates

The Illinois whitelist approach explicitly includes algae, while not listing any algae species as 

approved. Therefore all 27 species of nonindigenous algae are restricted by Illinois – including 

Actinocyclus normanii fo subsalsa, Bangia atropurpurea, Chaetoceros muelleri, Chroodactylon 

ornatum, Controcriba guillardii, Cyclotella atomus, Cyclotella cryptica, Cylindrospermopsis 

raciborskii, Diatoma ehrenbergii, Discostella pseudostelligera, Discostella wolterecki, 

Hymenomonas roseola, Nitellopsis obtusa, Pleurosira laevis, Skeletonema potamos, Skeletonema 

subsalsum, Sphacelaria fluviatilis, Sphacelaria lacustris, Stephanodiscus binderanus, 

Stephanodiscus subtilis, Thalassiosira baltica, Thalassiosira lacustris, Thalassiosira 

pseudonana, Thalassiosira weissflogii, Ulva flexuosa, Ulva intestinalis, and Ulva prolifera.

Illinois also includes aquatic invertebrates in its whitelist approach. We interpret this to include 

restrictions on the free-living annelids (Branchiura sowerbyi, Gianius aquaedulcis, Potamothrix 

bedoti, Potamothrix moldaviensis, Potamothrix vejdovskyi, and Ripistes parasita), Flatworms 

(Dugesia polychroa), insects (Acentria ephemerella and Tanysphyrus lemae), bryozoan 

(Lophopodella carteri), hydrozoa (Cordylophora caspia and Craspedacusta sowerbyi), and 

Amoebae (Psammonobiotus communis, Psammonobiotus dziwnowi, and Psammonobiotus 

linearis). 
34

3.4 Unregulated Species

Sixty four percent (116 of 181) of nonindigenous species established in the Great Lakes are not 

mentioned in any state or federal regulation. Seventy-eight of these species (67%) appear to be 

captured under Illinois’ whitelist approach; highlighting the strength of this approach in 

regulating additional species. This leaves 38 species (20%) entirely unregulated (Table 7).

Free-living micro-plankton and small benthic invertebrates are disproportionally represented 

among the nonindigenous species not captured by name but potentially regulated under the 

whitelist approach. There are no federal or state regulations which explicitly target any of the 27 

species of algae, eight free-living copepods, six annelids, three testate amoebae, two insects, two 

amphipods, one freshwater bryozoan, two freshwater hydroids, or one free-living flatworm 

which number among the Great Lakes nonindigenous species. Additionally, three of the five 

nonindigenous waterfleas (60%), 12 of 18 mollusks (66%), 28 of 56 plants (50%), and 16 of 21 

fish parasites/diseases (80%) are not mentioned by name. In contrast, most established 

nonindigenous fishes are identified in federal and state regulations; only 6 of the 27 

nonindigenous fishes (22%) are not regulated and half of those are captured by the whitelist 

approach leaving only 3 nonindigenous fish species entirely unregulated.

Table 7. Unregulated nonindigenous species. 

Fishes

 Esox niger

 Ictiobus cyprinellus

 Phenacobius mirabilis

Fish Parasites and Diseases

 Argulus japonicus

 Neoergasilis japonicus

 Salmincola lotae

 Bothriocephalus acheilognathi

 Dactylogyrus amphibothrium

 Dactylogyrus hemiamphibothrium

 Ichthyocotylurus pileatus

 Neascus brevicaudatus

 Scolex pleuronectis

 Timoniella sp.

 Acineta nitocrae

 Glugea hertwigi

 Heterosporis sp.

 Sphaeromyxa sevastopoli

 Trypanosoma acerinae

 Piscirickettsia cf. salmonis

Mollusks

 Bithynia tentaculata

 Viviparus georgianus

Plants

 Agrostis gigantea

 Impatiens glandulifera

 Lupinus polyphyllus

 Lycopus asper

 Mentha aquatica 

 Mentha gracilis

 Mentha spicata

 Myosotis scorpiodes

 Myosoton aquaticum

 Poa trivialis

 Polygonum persicaria

 Puccinellia distans

 Rumex longifolius

 Rumex obtusifolius

 Salix alba

 Salix fragilis

 Salix purpurea
35

Unregulated species are not necessarily innocuous. Icthyocotylurus pileatus, Heterosporis sp., 

and Bithynia tentaculata were rated in our previous assessments (GLERL TM-161) as having 

high environmental impact while Polygonum persicaria has high socioeconomic impact, 

Piscirikettsia cf. salmonis has moderate environmental impact and Salix fragilis has moderate 

socioeconomic impact – 6 out of 35 (17%) of unregulated species are thus still responsible for 

significant impacts.

4. Nonindigenous Species Control

In this section we summarize the control technologies that are available for control of various 

nonindigenous taxa. Information is drawn from the scientific literature and case studies. Where a 

particular method has been tried and proven ineffective, this is noted. Absence of information on 

the effect of a particular method on a particular species indicates only that we were unable to 

find information, not that the method has proven ineffective.

4.1 Algae

Options for control of established nonindigenous algae in open waters are limited. Only the 

largest two species (Ulva prolifera and Nitellopsis obtusa) can be effectively harvested 

mechanically, even then, they are often the first to recolonize (Pullman and Crawford 2010). 

Some species, particularly Cylindrospermopsis raciborskii, are associated with stratified water 

columns and mechanical destratification (mixing) may help prevent bloom development 

(Antenucci et al. 2005).

Nitellopsis obtusa is very sensitive to common algaecides containing copper and endothall based 

compounds. When N. obtusa is still low growing, algaecide can treat the entire organism. 

However, in taller individuals, the algaecide is absorbed in the top of the plant, killing that 

portion but leaving the bottom of the plant alive. This type of treatment has been found to be 

somewhat successful and is called a “hair cut treatment” by managers. The timing of the 

algaecide treatment is also important. Treatment early in the spring could help open up spawning 

habitat for native fish species, but N. obtusa or other nonnative aquatic plants are likely to 

recolonize these areas in the early summer. Algicides are generally less effective on planktonic 

algae, except for limited applications in smaller enclosed ponds. Note, the use of copper-based 

algicides may inhibit the degradation of cylindrospermopsin (Smith and Alexander 2008) and so 

are not a good choice when dealing with Cylindrospermopsis raciborskii.

Many of the invasive algae proliferate only in high-nutrient (A. normanii fo. subsalsa, Cyclotella 

atomus, Diatoma ehrenbergii, Discostella pseudostelligera, Stephanodiscus binderanus, 

Thalassiosira baltica, and Thalassioria pseudonana) or high-saline (Cyclotella cryptica, 

Thalassiosira lacustris, and Thalassiosira weissflogii) conditions or both (Bangia atropurpurea, 

Chaetoceros muelleri var. subsalsum, Contricribra guillardii, Stephanodiscus subtilis, Ulva 

flexuosa, Ulva intestinalis, and Ulva prolifera). Measures to reduce nutrient and salt pollution are 

usually the most effective options to control these species.
36

Management of Cylindrospermopsis raciborskii often focuses on alleviating impacts due to the

toxins that it produces. Potential management tools include absorption by activated carbon at 

point of use (Westrick et al. 2010), bacterial degradation (Donovan et al. 2008, Ho et al. 2012), 

and inactivation by chlorine, ozone, and/or hydroxyl radical treatments (Westrick et al. 2010). 

There are no known control methods for some planktonic algae, particularly those that are not 

associated with high-nutrient concentrations or salinity. Fortunately, these species 

(Chroodactylon ornatum, Hymenomonas roseola, Pleurosira laevis, Skeletonema potamos, 

Skeletonema subsalsum, Sphacelaria fluviatilis, and Sphacelaria lacustris) also tend to have low 

impact.

4.2 Plants

Although in some cases control options are extremely limited (e.g., to hand pulling) and large, 

established infestations may be difficult to control, control options are at least available for all 

the established nonindigenous plants. Many tools are available for control of aquatic plants, 

though rarely is any one method completely effective in eradicating a particular species. Several 

species of nonindigenous plants produce toxins that cause skin irritation, contact dermatis, 

allergic reactions in sensitive individuals, or even chemical burns (Pitcher 2004, Cooper and 

Johnson 1984, King County 2010). Proper protective clothing should be used when handling or 

burning the following: Conium maculatum, Iris pseudacorus, Polygonum persicaria, and 

Solanum dulcamara.

While small, new infestations of most invasive plants can be controlled by hand-pulling before 

seed-set, most invasive plants have characteristics (rhizomes, tubers, extensive roots, viable seed 

bank, etc.) which make this unfeasible for control of truly established or extensive infestations. 

Great care to remove all parts of the plant should be taken when implementing a physical method 

of control. To ensure removed plants will not sent out shoots, thoroughly dry all plant pieces 

(Jensen 2011).

Invasive tree species (Alnus glutinosa, Frangula alnus, Salix spp.) can be felled, but usually 

require application of an herbicide in order to prevent regrowth (USDA NRCS 2006).
37

Table 8. Physical control methods for invasive trees. Handpulling Felling/girdling + Herbicide Mowing/cutting/h arvesting

FloodingBurn

Alnus 

glutinosa

Effective

Frangula 

alnus

Plants 

<0.5cm

Effective if fruit 

are also 

removed/burned

Plants <2 years 

old. Most 

effective in 

spring.

Effective Repeat 

every 

2-3 

years

Salix spp. Effective 

for small 

seedlings

May need to be 

repeated, for 

large trees need 

to have the 

roots removed 

with machinery

Effective for 

seedlings

Mowing, cutting, or harvesting is frequently employed as a physical control for invasive plants. 

For most species, mowing should be conducted before seedset. Repeated mowing (often over 

several years) is recommended for species with extensive roots/rhizomes (Butomus umbellatus, 

Carex spp, Glyceria maxima, and Iris pseudacorus) to deplete those energy reserves (Jensen 

2011). Species which reproduce easily by fragmentation (e.g., Lysimachia spp. and

Myriophyllum spicatum) may be spread or promoted by attempts to harvest (Kennay and Fell 

2011). In other species, increased mowing promotes germination of the seed bank (Lythrum 

salicaria), aids seedling development (Rumex spp.), stimulates growth (Echinochloa crus-galli

and Juncus spp.), or increases competitive capacity of the invasive over natives (Cirsium 

palustre and Potamogeton crispus). 

Plants whose seeds require light to germinate (Echinochloa crus-galli and Lupinus polyphyllus)

may be controlled by seed burial via tilling or mulching (Cornell University 2012). Tilling may 

also be effective to kill seedlings (Alopecurus geniculatus and Rumex spp.).

Water level manipulation is frequently employed to control invasive plants (Toogood et al 2008, 

Lenseen et al 2000). Some species are sensitive to drying (Alopecurus geniculatus, Cabomba 

caroliniana, Hydrocharis morsus-ranae, Myriophyllum spicatum, Poa trivialis, and

Potamogeton crispus). Other species are sensitive to flooding (Epilobium hirsutum, Frangula 

alnus, Lythrum salicaria, and Typha angustifolia). Note that the length of time necessary and the 

sensitive season may vary with each species.
38

Controlled burns (Hanson et al. 2012, DiThomaso 2013) may be effective for some species 

(Frangula alnus, Lysimachia nummularia, Polygonum persicaria, and Typha angustifolia), but 

promote germination/regrowth in others (Iris pseudacorus and Lupinus polyphyllus).

Though usually limited to controlling small areas such as around docks or at swimming beaches, 

use of plastic or geotextile barriers or shading of the sediments (Forest Health Staff 2006) has 

proven effective for several species (Glyceria maxima, Myriophyllum spicatum, Najas minor, 

Polygonum persicaria, Potamogeton crispus, and Solanum dulcamara). 

Table 9. Physical control methods for invasive floating plants. Handpulling Mowing/cut ting/harvesti ng

Drying/drawdowns

Hydrocharis 

morsus-ranae

May provide 

temporary control

After turions have 

germinated but 

before extensive 

growth

Nasturtium 

officinale

Effective for small 

populations

Nymphoides 

peltata

Effective if all plant 

pieces are removed

Even with multiple 

harvests per year, 

complete control is 

unlikely.

Trapa natans roots easily uplifted 

and removed in small 

populations - all 

fragments must be 

removed

necessray for larger 

populations, 

fragments will 

regrow
39

Table 10. Physical control methods for invasive submerged plants. Handpulling Mowing/cuttin g/harvesting Seed burial/Tilling/ mulch Drying/

drawdownsShading/Barriers

Cabomba 

caroliniana

avoid late 

season 

when the 

plant is 

brittle

likely to 

provide relief 

only for a few 

weeks

Will reduce 

growth but 

rootball must 

be dried 

completely 

or it will 

return

Myriophyllum 

spicatum

Effective 

for small 

populations

Only after all 

niches filled -

otherwise this 

will enhance 

the rate of 

spread. 

Multiple 

harvest per 

season

Most 

effective if 

plants 

exposed to 

several 

weeks drying 

and root 

crowns 

exposed to 

sub-freezing 

temperatures

Cover 

sediment 

with 

opaque 

fabric

Najas minor Physical 

removal 

provides only 

short-term 

relief

Benthic 

barriers 

may be 

effective

Potamogeton 

crispus

Effective if 

crowns, 

turions and 

fragments 

are 

removed

Use only if 

monoculture

Use only if 

monoculture

Autumn 

drawdown 

will kill 

turions

Effective
40

Table 11. Physical control methods for invasive emergent monocot species. Handpulling Mowing/cutting/harv esting Seed burial/Tilling/ mulch Drying/ drawdowns Flooding

BurnShading/Barriers

Agrostis 

gigantea

Effective 

for small 

populations

<3 inches

Alopecurus 

geniculatus

effective 

only if seed 

bank not 

extensive

Effective Not 

effective

Butomus 

umbellatus

rhizomes 

must be 

removed

multiple 

cuts per 

year will 

reduce 

abundance 

and 

prevent 

spread

Not 

effective

Carex spp. rhizomes 

must be 

removed

>7cm Not 

effective

Not 

effective

Echinochloa 

crus-galli

Not 

effective

<1 cm 

prevents 

germination

Glyceria 

maxima

Effective if 

rhizomes 

can be 

removed

2-3 times 

per 

summer 

may 

deplete 

roots and 

rhizomes

May be 

used 

together 

with other 

methods

Cover 

with 

black 

plastic 

for 5-

6 

weeks

Iris 

pseudacorus

Rhizomes 

must be 

removed.

Remove 

leaves and 

stems 

above 

water level 

before 

flowing. 

May need 

to be 

repeated 3-

4 years.

Not 

recommende

d as this 

encourages 

reprouting 

from 

rhizomes

Juncus spp. This may 

increase 

Not 

effective

Not 

effective
41

the seed 

bank as 

well as 

promote 

regrowth.

Poa trivialis This 

species 

has a low 

tolerance 

for 

drought.

Sparganium 

glomeratum

roots are 

easily lifted 

from 

sediment, 

but must be 

removed

roots are 

easily 

lifted from 

sediment, 

but must 

be 

removed

Typha 

angustifolia

during the 

growing 

season and 

again just 

before 

flowers 

reach 

maturity

Flooding 

can be 

effective if 

it triggers 

anaerobic 

respiration 

in the 

sediments, 

or 

following 

cutting

May be 

effective if 

repeated 

several times 

or rhizomes 

are exposed
42

Table 12. Physical control methods for invasive emergent broadleaf plants. Handpulling Mowing/cuttin g/harvesting Seed burial/Tilling/ mulch Flooding Burn

Shading/Barriers

Chenopodium 

glaucum

spring 

and fall

Cirsium 

palustre

cut just 

below 

surface 

before 

flowering

Repeated 

(>3x per 

year) close 

mowing for 

3-4 years 

necessary

Conium 

maculatum

Taproot 

must be 

removed

Mow before 

flowering

Epilobium 

hirsutum

Effective if 

rhizomes 

can be 

removed

>18 weeks 

can reduce 

roots and 

leave the 

plant more 

susceptible 

to other 

control 

methods

Impatiens 

glandulifera

Before seedsetCut below 

the lowest 

node to 

prevent regrowth. 

Stems left 

must be 

crushed to 

prevent 

regrowth.

Lupinus 

polyphyllus

Root system 

should be 

severed 

below the 

crown.

Generally 

not effective 

unless very 

frequent

Not 

effective, 

promotes 

germination

Lysimachia 

nummularia

All 

fragments 

and 

rhizomes 

May contain 

the existing 

population, 

but will not 
43

must be 

removed to 

prevent 

resprouting

eradicate it.

Lythrum 

salicaria

Remove 

entire root 

system 

before 

seedset 

using a 

cultivator

Only if 

followed by 

flooding, 

otherwise it 

promotes 

germination

> 30cm 

for over 7 

weeks, or 

following 

mowing

Not 

effective

Mentha spp. Effective for 

small 

populations

Soil 

barriers 

(edging) 

may 

contain 

spread via 

rhizomes

Myosoton 

aquaticum

Before seeds 

form

Polygonum 

persicaria

Effective Effective to 

kill seeds

Solarizing 

black 

plastic 

will kill

seeds

Rorippa 

sylvestris

Small 

rhizome 

fragments 

are difficult 

to remove

Rumex spp. To prevent 

seedset, but 

may aid 

seedling 

development 

and promote 

branching

Repeated 

will 

eliminate 

seedlings

Solanum 

dulcamara

all parts of 

roots must 

be removed

Not effective 

unless 

followed by 

root removal

Cover 

with 

geotextile 

cloth for 

2 years

Solidago 

sempervirens

Deadheading 

prior to 

seedset is 

effective

Deadheading 

prior to 

seedset is 

effective
44

Many chemical herbicides are available for the control of invasive plants. However, only a 

handful of these are approved for use in open waters. Thus while the following notes herbicides 

and chemical families that have proven effective for particular species, options in open water are 

MUCH more limited than this suggests.

Glyphosate is an EPSP Synthase inhibitor (Herbicide Family 9) which leads to the depletion of 

the aromatic amino acids tryptophan, tyrosine, and phenylalanine (Shaner 2006). This systemic 

broad-spectrum herbicide is used to control floating-leaved plants like water lilies and shoreline 

plants like purple loosestrife. It is generally applied as a liquid to the leaves. Glyphosate does not 

work on underwater plants such as Eurasian watermilfoil. Glyphosate is non-selective and will 

injure any plant tissue with which it comes in contact. It is toxic to frogs and tadpoles. It is 

commonly used as a foliar spray and proven effective against Alnus glutinosa (stump 

application), Carex spp., Cirsium palustre (applied to cut stems), Conium maculatum, Frangula 

alnus (applied to basal stems or foliar), Glyceria maxima, Ornamental jewelweed (Impatiens 

glandulifera), Iris pseudacorus, Juncus spp., Lysimachia spp., Lythrum salicaria, Mentha spp., 

Giant chickweed (Myosoton aquaticum), dock (Rumex) spp. (at early heading), Willow (Salix)

spp., and Typha angustifolia. It is effective against Echinochloa crus-galli if applied to plants <5

cm in height. Because of its non-selective nature, re-seeding (with natives capable of 

outcompeting seedlings) is required to prevent regrowth of species such as lupine (Lupinus 

polyphyllus) and Rough-stalked meadow grass (Poa trivialis). While not directly tested on 

invasive Alopecurus geniculatus, it has been proven effective against related species. It is not 

effective against Nasturtium officinale in flowing water. It will kill above-ground biomass of 

Great Hairy Willowherb (Epilobium hirsutum) but repeated applications are needed to weaken 

roots. Likewise it is marginally effective against Creeping Yellow Cress (Rorippa sylvestris) and 

Nymphoides peltata. Some applicators report control of Butomus umbellatus with glyphosate. 

Glyphosate-resistant strains of Redtop (Agrostis gigantea) and Lady’s Thumb (Polygonum 

persicaria) are becoming common. Plants can take several weeks to die and a repeat application 

is often necessary to remove plants that were missed during the first application.

ACCase (acetyl-Coenzyme A carboxylase) inhibitors (Herbicide Family 1) block the first step in 

fatty acid synthesis in grasses (Armstrong 2012). With a few specific exceptions, herbicides in 

this family are not approved for use in aquatic environments, but can be used to control these 

invasives in upland settings. Most of the herbicides in this family are most effective against 

seedlings when applied early post-emergence. Broadleaf plants have a natural resistance to 

herbicides in this family. Some herbicides in this family control only specific subsets of grasses. 

Cyhalofop-butyl is highly toxic to both freshwater and estuarine/marine animals on an acute 

exposure basis. Data indicate that the major degradation products for cyhalofop-butyl are nontoxic to most aquatic organisms at normal application levels; however for endangered species of 

estuarine/marine fish, estuarine/marine invertebrates, and freshwater fish this low toxicity is still 

considered a concern and use prohibited where such species are of concern. It is prohibited for 
45

aquatic use except in rice fields. However, most varieties of rice are now tolerant of cyhalofopbutyl. Clethodim is slightly toxic to fish and aquatic invertebrate species. Clethodim may be 

highly persistent in the aquatic environment. It can be used to control invasive plants in upland 

areas, but not for direct aquatic application. Sethoxydim will not harm broadleaf herbs, sedges or 

woody plants. It is only mildly toxic to aquatic animals, but degrades rapidly (<1 hour) in open 

water. Dichlofop, fluazifop and haloxyfop are very toxic to aquatic organisms, may cause longterm adverse effects in the aquatic environment. Fluazifop-p cannot be used in Forest 

Sustainability Certified Areas. Fluazifop controls Water Foxtail (Alopecurus geniculatus).

Table 13. Plants susceptible to ACCase inhibitors. 

Cyhalofop Clethodium Dichlofop Fluazifop Haloxyfop Sethoxydim

Agrostis 

gigantea

Proven for 

related 

species

Alopecurus 

geniculatus

Effective Effective

Echinochloa 

crus-galli

Effective Effective Effective Effective Effective Effective

Frangula 

alnus

Effective

Juncus spp. Effective

ALS (acetolactate synthase) inhibitors (Herbicide Family 2) block the production of branched 

chain amino acids (isoleucine, leucine and valine). Herbicide resistance is common in this family

(Armstrong 2012). Corn is often susceptible to herbicides in this family thus many are not 

recommended for use near croplands. Rimsulfuron and DPX-79406 (1:1 premix of nicosulfuron 

and rimsulfuron) are registered in Ontario and can be used with corn. Bispyribac is registered for 

use in ricefields. Rimsulfuron is most effective in dry conditions. Chlorsulfuron should not be 

used on powdery, dry, light or sandy soils. Imazapyr is a systemic, broad-spectrum, slow-acting 

herbicide. When applied as a liquid it is used to control emergent plants like spartina, reed 

canarygrass, phragmites, and floating-leaved plants like water lilies but does not work on 

underwater plants such as Eurasian Watermilfoil. Imazapyr is effective on some sedge species 

and most effective when water levels are low and during calm weather. Imazapic is similar, but 

non-selective.
46

Table 14. Plants susceptible to ALS inhibitors. bensulfuron methyl

BispyribacchlorsulfuronImazapyrImazamox/imazapic/imazaquinmetsulfuronpenoxsulamrimsulfuronSulfometuronsulfosulfuron

Agrostis 

gigantea

proven 

effective 

on 

related

species

effective 

if kept 

dry

Butomus 

umbellatus

effective 

during 

calm 

weather

Carex spp. proven 

effective on 

some sedge 

species

Cirsium 

palustre

effective 

foliar

Conium 

maculatum

effective effective effective

Echinochloa 

crus-galli

effective 

postemergent

effective effective effective effective

Frangula 

alnus

effective 

foliar

effective 

foliar

Glyceria 

maxima

effective 

only when 

water 

levels are 

low

Hydrocharis 

morsusranae

effective effective effective 

Impatiens 

glandulifera

effective 

Iris 

pseudacorus

excellent 

for large 

infestations

Juncus spp. effective 

Lupinus 

polyphyllus

effective effective

Lysimachia 

vulgaris

effective
47

Lythrum 

salicaria

effective

Marsilea 

quadrifolia

used in 

Japan

Poa trivialis currently 

being 

labelled 

for 

selective 

control in 

creeping 

bentgrass

Rumex spp. effective 

on young 

plants

effective

Salix spp. effective 

on fully 

leavedout 

plants

Typha 

angustifolia

effective
48

Microtubule protein inhibitors (Herbicide Family 3) interfere with the alignment of the spindle 

apparatus during mitosis and prevent normal cell division in root tissue; thus they inhibit root 

growth (Armstrong 2012). Pendamethalin is used for post-emergent control of Echinochloa crusgalli and trifluralin for control of Rorippa sylvestris. Pendamethalin cannot be used in Forest 

Sustainability Certified Areas or open waters.

Synthetic auxins (Herbicide Family 4) mimic the natural plant hormone IAA (indole-3-acetic 

acid). These herbicides affect cell wall plasticity and nucleic acid metabolism which leads to 

inhibited cell division and growth in the meristem (Armstrong 2012). These herbicides are 

generally selective for broadleaf control. 2,4-D is approved in both granular and liquid forms; by 

court-order the butoxy-ethyl-ester formulation of 2,4-D cannot be used in waters with threatened 

and endangered salmon-bearing waters in the Pacific Northwest. Picloram and Triclopyr have 

been proven effective in control of Conium maculatum, Frangula alnus, and Rumex spp. 

Triclopyr also is effective for control of Alnus glutinosa, Impatiens glandulifera, Lysimachia 

vulgaris, Lythrum salicaria, Myriophyllum spicatum, Salix spp., Solanum dulcamara, and Trapa 

natans. There are two formulations of triclopyr. Triclopyr should be used in its amine form in 

wetlands. The TEA formation of triclopyr is registered for use in aquatic or riparian 

environments. Many native aquatic species are unaffected by triclopyr; it is very useful for 

Purple Loosestrife control since native grasses and sedges are unaffected by this herbicide.

Photosystem I inhibitors (Herbicide Family 22) capture electrons from photosystem I and are 

reduced to form free radicals that destroy cell membranes (Armstrong 2012). Diquat is a fastacting, non-selective contact herbicide which destroys the vegetative part of the plant but does 

not kill the roots. It is applied as a liquid. Typically diquat is used primarily for short term (one 

season) control of a variety of submersed aquatic plants. It is very fast-acting and is suitable for 

spot treatment. However, turbid water or dense algal blooms can interfere with its effectiveness. 

Diquat provides effective control of Hydrocharis morsus-ranae, Myriophyllum spicatum, Najas 

minor, and Typha angustifolia. It can be used to control Potamogeton crispus if applied before 

turion production and has been proven in laboratory tests to be effective against Cabomba 

caroliniana (Michigan applicators report it to be ineffective against Cabomba in the field) . 

Some applicators report control of submerged Butomus umbellatus with diquat in sites that have 

limited dilution potential. Paraquat is effective against Echinochloa crus-galli when applied to 

plants less than five centimeters in height. 

Photosystem II Inhibitors (Herbicide Family 5-7) inhibit photosynthesis by binding to the QB‐

binding niche on the D1 protein of the photosystem II complex in the chloroplast (Armstrong 

2012). It blocks electron flow from QA to QB and stops carbon dioxide fixation and production

of ATP and NADPH2 which are needed for plant growth and development. Death occurs from

free radicals destroying cell membranes. Herbicide resistance is increasingly common in this 

group. Herbicides in this class have proven effective in control of Agrostis gigantea, Conium 

maculatum, Myriophyllum spicatum, and Potamogeton crispus. This family of herbicides is most 
49

effective against Echinochloa crus-galli when used post-emergence on plants <5 cm. 

Hexazinone and simazine cannot be used in Forest Sustainability Certified Areas. 

Table 15. Nonindigenous plants susceptible to Photosystem II inhibitors. Atrazine Hexazinone Linuron Metribuzin Simazine Terbacil Terbutryn

PropanilButhidazole

Agrostis 

gigantea

Effective

Conium 

maculatum

Effective Effective Effective

Echinochloa 

crus-galli

Effective Effective Effective Effective Effective Effective Effective

Myriophyllum 

spicatum

Effective

Potamogeton 

crispus

Effective

Pigment synthesis inhibitors (Herbicide Family 12, 13 and 27) are collectively called ‘bleachers’ 

because of the characteristic whitening of plant tissue (Armstrong 2012). These may inhibit 

chlorophyll or other pigments such as carotenoids. Fluridone is a slow-acting systemic herbicide 

used to control Eurasian Watermilfoil and other underwater plants. It may be applied as a pellet 

or as a liquid. Fluridone can show good control of submersed plants where there is little water 

movement and an extended time for the treatment. Its use is most applicable to whole-lake or 

isolated bay treatments where dilution can be minimized. It is not effective for spot treatments of 

areas less than five acres. It is slow-acting and may take 6-12 weeks before the dying plants fall 

to the sediment and decompose. Granular formulations of fluridone are proving to be effective 

when treating areas of higher water exchange or when applicators need to maintain low levels 

over long time periods. Although fluridone is considered to be a broad spectrum herbicide, when 

used at very low concentrations, it can be used to selectively remove Eurasian Watermilfoil. 

Some native aquatic plants, especially pondweeds, are minimally affected by low concentrations 

of fluridone. Cabomba caroliniana has been controlled in one waterbody in Michigan by whole 

lake application of fluridone (20 ppb). 
50

Table 16. Nonindigenous plants susceptible to pigment synthesis inhibitors. 

Fluridone Norflurazon Isoxaflutole Mesotrione

Agrostis 

gigantea

Proven effective 

on related 

species

Proven effective 

on related 

species

Cabomba 

caroliniana

Effective

Echinochloa 

crus-galli

Effective

Myriophyllum 

spicatum

Effective

Najas marina Effective

Najas minor Effective

Potamogeton 

crispus

Effective if 

applied before 

dieback

PPO Inhibitors (Herbicide Family 14) inhibit the photoporphyringogen oxidase, an enzyme that 

is responsible for chlorophyll and heme biosynthesis (Armstrong 2012). PPO inhibition leads to 

accumulation of PPIX (protoporphyrin IX), which creates free radical oxygen in the cell and 

destroys cell membranes. Carfentrazone is proven effective for control of Myriophyllum 

spicatum. Flumioxazin has proven effective against Cabomba caroliniana in laboratory tests and 

in many Michigan waterbodies (200 ppb flumioxazin). LH Some applicators report control of 

Nitellopsis obtusa with 150 ppb flumioxazin.

VLFA Inhibitors (Herbicide Family 15) are herbicides that inhibit very long chain fatty acid 

synthesis in shoot tissue during germination of sensitive plants. Napropamide can be used for 

pre-emergent control of Alnus glutinosa.

Dichlobenil (Herbicide Family 20) inhibits actively dividing cells by disrupting formation of the 

cell plate. Germinating seedlings and actively growing plants are most affected by dichlobenil. 

Dichlobenil is effective against Myriophyllum spicatum, Nymphoides peltata, Potamogeton 

crispus, and Rorippa sylvestris. In rotation, it can be effective against Polygonum persicaria (this 

plant is known for developing resistance). 

Endothall’s herbicidal mode of action is not clear. It interferes with plant protein synthesis in 

some way and also affects lipid synthesis and dipeptide and proteinase activities (McDonald et 

al. 1993). The net result is a wilting of leaf tissue. Endothall may be applied in a granular or 

liquid form. Typically endothall compounds are used primarily for short-term (one season) 

control of a variety of aquatic plants. However, there has been some recent research that 

indicates that when used in low concentrations, endothall can be used to selectively remove 
51

exotic weeds, leaving some native species unaffected. Because it is fast acting, endothall can be 

used to treat smaller areas effectively. Endothall has been proven effective against Cabomba 

caroliniana (Michigan applicators have reported that Cabomba populations in Michigan do not 

appear to be controlled by endothall in the field), Myriophyllum spicatum, and Najas spp. It can 

be effective against Potamogeton crispus if used before turion production. 

Alkali grasses, such as Puccinellia distans, and other salt-tolerant invasives, such as Solidago 

sempervirens, outcompete other grasses in most brackish conditions. Thus, management efforts 

in the freshwater Great Lakes regions targeted at reducing salt contamination may benefit efforts 

to control these species.

Biological controls are employed for some plant species. Biological control agents do not 

completely eliminate the target, but when successful, can suppress populations to a 

nonsignificant level (Rees et al. 1996). 

Grass Carp (Ctenopharyngodon idella) are voracious consumers of aquatic plants and frequently 

employed in the southern states for control of invasive plants. However, diploid (fertile) Grass 

Carp are illegal for use in most Great Lakes states. Grass Carp are emerging as an “aquatic 

species of concern” for the Great Lakes region (Great Lakes ANS Panel, Research Coordination 

Committee). The use of certified triploid (sterile) Grass Carp is allowed in Illinois, New York,

and Pennsylvania, with the correct permits (Shiels and Hartle 2012, NYSDEC 2013). Since 

1963, the Grass Carp, Ctenopharyngodon idella has been released to suppress Eurasian 

Watermilfoil and other nuisance aquatic plants in numerous sites within North America (Julien 

and Griffiths 1998, CEH 2004d). It has been found that Grass Carp may only eat Eurasian 

Watermilfoil after native plants have been consumed (IL DNR 2009a). To achieve control of 

Eurasian Watermilfoil generally means the total removal of more palatable native aquatic species 

before the Grass Carp will consume Eurasian Watermilfoil. In situations where Eurasian 

Watermilfoil is the only aquatic plant species in the lake, this may be acceptable. However, 

generally Grass Carp are not recommended for Eurasian Watermilfoil control. Likewise, Grass 

Carp will provide effective control of Potamogeton crispus, but may feed on native plants (CEH 

2004e). Other bottom feeding fish, such as Common Carp, do not feed on P. crispus, but they 

create turbid water conditions and may prevent the growth of this plant species (CEH 2004e).

Grass Carp will eat Cabomba caroliniana, but it is not their preferred food source. Grass Carp 

also feed on Hydrocharis morsus-ranae. 

In their native range in Europe, ducks have been known to graze extensively on Butomus 

umbellatus (Hroudová et al. 1996). Beaver can have huge impact on willow species and are 

capable of eradicating entire stands (USDA NRCS 2002). Muskrat (Ondatra zibethicus) 

populations can have a serious impact on Typha populations. However, large populations of 

muskrats can shift to other plants species and have a long-term detrimental effect on the 

vegetation community (Miklovic 2000).
52

Van Leeuwen (1983) found that a combination of European grazers (rabbits, the hoverfly 

Cheilosia grossa, and Epiblema scutula) resulted in an approximately 30% reduction in flower 

heads on Cirsium palustre. Furthermore, plants that had suffered predation had a reduced stem 

height, resulting in a reduced seed dispersal distance of surviving achenes (van Leeuwen 1983). 

Additional research is needed to determine if native species of rabbits and insects could have 

similar results on controlling C. palustre in the Great Lakes. Bitter Dock (Rumex obtusifolius) is 

avoided by rabbits, but docks appear to be a favorite food plant of deer (Amphlett and Rea 

1909). 

Grazing of domestic herbivores can be an effective method of control for many palatable species. 

Allowing cattle or sheep access to areas infested with Impatiens glandulifera will control the 

population and the spread of the species either by direct grazing or by trampling of young 

seedlings (CEH 2004b). Cattle, horses, and sheep graze on Juncus spp., but the extent of control 

gained from grazing is unknown (CEH 2004a, Cosyns et al. 2005). Docks (Rumex spp.) are 

grazed by cattle, sheep, and goats but not by horses. Targetted grazing by sheep has been used as 

a biocontrol for Purple Loosestrife (Kleppel and LaBarge 2011). Goats are attracted to the 

flowering stage of many thistles, including Cirsium palustre. Only about 0.5% of thistle seeds 

that pass through their digestive systems remain viable, making it unlikely that they would aid in 

the spread of this species. Effective grazing could reduce Marsh Thistle populations, although it 

is unclear whether grazing would ultimately control C. palustre via the trampling of rosettes or 

facilitate its spread through the creation of safe sites for germination (Fraser 2000). Reseeding of 

native vegetation may enhance the success of prior control efforts. Moreover, goats do not select 

for Marsh Thistle and may also eat native thistles in intermingled communities (Popay and Field 

1996). Heavy grazing will eliminate Typha spp. from riparian corridors; however, this technique 

might also affect other native species (Stevens and Hoag 2006).

Five species of beetles have been approved for the biocontrol of Lythrum salicaria (Blossey et al.

1994ab). Galerucella calmariensis and G. pusilla are both leaf-feeding chrysomelids. These 

beetles defoliate and attack the terminal bud area, drastically reducing seed production. The 

mortality rate to Purple Loosestrife seedlings is high. Evidence of Galerucella spp. damage are 

round holes in the leaves. Four to six eggs are laid on the stems, axils, or leaf underside. The 

larvae feed constantly on the leaf underside, leaving only the thin cuticle layer on the top of the 

leaf. Initial introductions in eastern North America occurred in Virginia, Maryland, 

Pennsylvania, New York, Minnesota, and southern Ontario in August 1992 (Hight et al. 1995). 

In 1992 these three beetles were released in Washington. Galerucella spp. populations visibly 

impacted Purple Loosestrife stands by 1996 (Washington State Department of Ecology 2012). In 

the Great Lakes region, Sea Grant conducted an extensive, multi-state program involving youth 

in raising and releasing Galerucella beetles for control of L. salicaria (Michigan Sea Grant 

2001). 

Hylobius transversovittatus is a root-mining weevil that also eats leaves (Wilson et al. 2004). 

This beetle eats from the leaf margins, working inward. The female crawls to the lower 2-3 
53

inches of the stem then bores a hole to the pithy area of the stem where 1-3 eggs are laid daily

from July to September. Or, the female will dig through the soil to the root, and lay eggs in the 

soil near the root. The larvae then work their way to the root. H. transversovittatus damage is 

done when xylem and phloem tissue are severed, and the carbohydrate reserves in the root are 

depleted. Plant size is greatly reduced because of these depleted energy reserves in the root. The 

presence of larvae is evidenced by zig-zag patterns in the root.

Nanophyes marmoratus and N. brevis are seed eating beetles (Blossey 2002a, Wilson et al.

2004). Young adults feed on shoot tips, later feeding on the flowers and closed flower buds. 

Sixty to one hundred eggs are laid in the immature flower bud. Seed production is reduced by 

60%. There were two test sites releases in 1996. Approval to introduce N. marmoratus was 

granted followed by introductions in New York and Minnesota in 1994. Additional releases 

occurred in New Jersey in 1996. N. marmoratus has also been released in Ohio (Ohio EPA 

2001). Release of N. brevis planned for 1994 was delayed due to contamination of the original 

shipment with a parasitic nematode (Piper 1997). This infection appeared benign for N. brevis, 

however, due to the potential for non-target effects of the nematode after introduction into North 

America, only disease free specimens should be introduced, which, at present, effectively 

precludes the introduction of N. brevis (Blossey 2002a). Bayeriola salicariae, a gall midge, was 

studied and screened between 1990 and 1992 (Blossey and Schroeder 1995). Based on results 

indicating a potential wider host range, the gall midge B. salicariae was not proposed for 

introduction (Blossey and Schroeder 1995).

Many invasive plants are not susceptible to grazing. For example, Lupines (Lupinus polyphyllus) 

may be toxic, and populations often increase in grazed (pasture) systems. Several native insects 

feed on Lupines, but are considered insufficient for control (DiTomaso 2013). Chemical defense 

indicates that the use of biocontrol agents on invasive populations of Cabomba caroliniana may 

not be a viable approach (Morrison and Hay 2011). When fed on by crayfish and snails, C. 

caroliniana induces a chemical defense mechanism to deter both herbivores and microbes that 

typically attack plants via openings left by herbivores (Morrison and Hay 2011).

Research into potential biocontrol via host-specific and native insects is ongoing for many 

species of invasive aquatic plants. A North American weevil, Euhrychiopsis lecotie, may be 

associated with natural declines of Myriophyllum spicatum at northern lakes (Sheldon 1994, 

Creed and Sheldon 1995). E. lecotei feeds on the new growth of M. spicatum and can help keep 

populations under control; it is common for the populations of E. lecotei and M. spicatum to 

exhibit the classic predator-prey cycles (Creed and Sheldon 1995, Michigan Sea Grant 2012b). 

Studies have found the herbivorous weevil causes significant damage to Eurasian water-milfoil 

while having little impact on native species, suggesting the insect as a potential biocontrol agent 

(Creed and Sheldon 1995). Female weevils have higher fecundity when raised on M. spicatum as 

opposed to native M. sibiricum (Solarz and Newman 1996, Creed 1998, TNC Vermont 1998, 

Sheldon and Jones 2001).
54

The defoliating Hemlock Moth (Agonopterix alstroemeriana) was accidentally introduced to the 

United States, but it is now being investigated as a potential biocontrol agent because of its 

monophagous (feeding on a single food source) association with Conium maculatum (Castells 

and Berenbaum 2006). These moth larvae feed on the young stem tissue, flowers, and seeds 

(Forest Health Staff 2006d). High densities of A. alstroemeriana have been effective drivers of 

plant mortality in C. maculatum stands in the western United States, where several hundred 

larvae have been reported from a single plant. However, as a chemical defense, alkaloid 

production appears to increase with A. alstromeriana herbivory, potentially driving surviving 

populations to higher levels of toxicity over time (Castells et al. 2005). Furthermore, A. 

alstroemeriana was found to be targeted by a predatory wasp (Euodynerus foraminatus) in 

Illinois, suggesting that the effectiveness of biocontrol may be lessened in the Midwest and other 

locations where E. foraminatus exerts top-down pressure on A. alstroemeriana (Castells and 

Berenbaum 2008). Although A. alstroemeriana is widespread in the United States, larvae may 

still be difficult to obtain for biocontrol purposes (Castells and Berenbaum 2006). Trichoplusia 

ni, the cabbage looper, is a generalist lepidopteran that is found throughout the United States and 

occasionally feeds on C. maculatum. Overall growth of T. ni is not stunted, but larvae raised on 

diets enriched with the piperidines found in C. maculatum develop slower. A prolonged larval 

stage makes T. ni more vulnerable to predators and could reduce overall biocontrol capabilities 

(Castells and Berenbaum 2008). Papulio poluxenes, black swallowtail butterfly, will lay eggs on 

C. maculatum, but Feeny et al. (1985) found low larvae survivorship in central New York.

While there are no specific biocontrol agents for Cirsium palustre (GLIFWC 2006), herbivory by 

a variety of species may be beneficial but requires additional research. Promising biocontrol 

candidates include a European seedhead fly, Terellia ruficauda (Fraser 2000, OLA, and MAFF 

2002); the seed-eating weevil, Rhinocyllus conicus, currently undergoing experimental trial in 

the Robson Valley Forest District, British Columbia (OLA and MAFF 2002, USDA Forest 

Service 2005b); and the glassy cutworm, Apamea devastator (native in New York and Ohio; 

Volger and Stressler 2011). The latter is an indiscriminate herbivore known to feed on C. 

palustre and may help control Marsh Thistle; however, this moth feeds on a broad spectrum of 

additional plants. Larvae of the artichoke plume moth (Platyptilia carduidactyla) also feed on 

Marsh Thistle, but as its common name suggests, this species is considered a pest to artichokes

(Winston et al. 2008). Furthermore, the moth’s native range is south of the Great Lakes. 

Occasionally Cheilosia corydon, a fly native to Italy, feeds on Marsh Thistle (Winston et al. 

2008). This fly was released in Oregon in 1991 to control several invasive thistle populations. 

However, since its release, C. corydon populations have attacked native and exotic thistles 

indiscriminately (ODA Plant Division 2011). 

Elephant moth (Deilephila elpenor) feeds on Epilobium hirsutum, but is not a native to the Great 

Lakes (Hoskins 2012, Pittaway 2012). Genetic material extracted from E. hirsutum individuals 

displaying phyllody of flowers and/or plant yellowing revealed infection by epilobium phyllody 
55

(EpPh) phytoplasma, an obligate, parasitic bacteria that attach to phloem tissue (Alminaite et al. 

2002). The ability of this phytoplasma to act as a biocontrol agent is still unknown.

The use of the stem-boring larvae of the weevils Apion violaceum and A. miniatum has been 

investigated for controlling Rumex obtusifolius (Hopkins 1980, Freese 1995). In the UK and 

elsewhere, the Chrysomelid Beetle (Gastrophysa viridula) has been investigated as a biocontrol 

agent for both R. obtusifolius and R. crispus (Bentley et al. 1980). Larvae of the leaf-mining fly 

Pegomya nigritarsis cause blotch mines on leaves of R. obtusifolius (Whittaker 1994). R. 

obtusifolius is the preferred host plant of Coreus marginatus and has been shown to moderately 

reduce its seed viability (Hruskova et al. 2005). 

In its native range in China, the leaf beetle Galerucella birmanica has significant negative 

impacts on Trapa natans populations (Ding et al. 2006). However, this species has many other 

host species in the United States, making it unsuitable for use as a biocontrol agent (Maryland 

Sea Grant 2012).

The species Aphis fabae, Impatientinum balsamines, and Deilephila elpenor are known to feed 

on Ornamental Jewelweed (Impatiens glandulifera), but their capacity to act as biological control 

agents is still unknown (Beerling and Perrins 1993). Although an initial experiment by Tanner 

(2011) indicated that D. elpenor exhibited lower biomass and survivorship when raised on I. 

glandulifera.

Aphids may occasionally feed on Juncus spp., but most rushes are fairly resilient to extensive 

damage from insect or diseases (Stevens and Hoag 2003). Gypsy moths can defoliate Purple 

Willow and willow midges can cause significant (though rarely fatal) damage. The native 

boring-moth larvae (Arzama spp.) have been reported to cause damage to Typha stands, but their 

use as a species-specific biological control is unknown (Miklovic 2000). Chrysolina herbacea

feeds on Mentha aquatica, despite the deterrents this species produces to minimize damage 

caused by herbivores (Atsbaha Zebelo et al. 2011). The fruit fly Acinia picturata has been known 

to use Pluchea odorata as a host, but it is unknown if this species could be used a biological 

control agent (Stegmaier 1967).

Pathogens are also being explored as potential biocontrol agents (biopesticides regulated by EPA 

as pesticides). Conium maculatum is susceptible to multiple viruses, including ring spot virus, 

carrot thin leaf thin virus (CTLV), alfalfa mosaic virus (AMV), and celery mosaic virus (CeMV) 

(Howell and Mink 1981). However, viral infections appear to stunt growth rather than cause 

mortality, diminishing their potential for biocontrol (Howell and Mink 1981, Pitcher 2004). 

Another disadvantage to using these types of biological control agents is the potential for them to 

escape into neighboring habitats, especially agricultural fields (J. McHenry pers. comm. in 

Pitcher 1985). The fungal pathogen Exserohilum monoceras has shown some success in 

controlling Barnyard Grass, Echinochloa crus-galli (Catindig et al. 2009). In its native range,

Impatiens glandulifera has been known to harbor Puccinia komarovii (a rust pathogen), which is 
56

currently undergoing research as a control agent (Tanner 2011). Laboratory research has shown 

that the fungus Mycoleptodiscus terrestris reduces the biomass of Myriophyllum spicatum

significantly and may be a possible biocontrol agent (IL DNR 2009a). Although Polygonum 

persicaria plants are susceptible to Arabis mosaic virus, no research has been undertaken on the 

development of biological control agents, whether viral or fungal.

The Leaf Spot Fungus Ramularia rubella causes red spots to develop on dock leaves but has no 

major effect on plant survival. The Rust Fungus Uromyces rumicis is also non-systemic but has 

been shown to have some potential as a biological control agent (Inman 1971, Schubiger et al. 

1986). Dock species are also an alternate host for number of viruses, fungus (Dal Bello and 

Carranza 1995), and nematodes (Townshend and Davidson 1962, Edwards and Taylor 1963).

Willow blight is a serious pest to Salix plantings, and has been documented to kill plants 

damaged by storms, indicating a potential for use in combination with mechanical control

(USDA NRCS 2002).

Manipulation of competitors can also be an important element of biological control. Control of 

many species has the best long-term efficacy when followed by replanting with native species,

which can outcompete seedlings of the invasive. For example, revegetation of disturbed riparian 

sites can be used to prevent Purple Loosestrife establishment and to reduce re-establishment after 

control procedures are applied. Fowl Mannagrass (Glyceria striata), Foxtail Sedge (Carex 

alopecoidea), and Reed Canarygrass (Phalaris arundinacea) have achieved dominance and 

prevented re-invasion in plots where Purple Loosestrife was experimentally removed (Lui et al.

2005). Smartweed (Polygonum lapathifolium) is reported to out-compete Purple Loosestrife

during its first year of growth. Seeding Japanese Millet (Echinocloa frumentacea, also called 

billion-dollar grass) after drawdown and before Purple Loosestrife seedlings began to grow 

provided control (Jacobs 2008). Combining herbicides with overseeding an alternative desired 

grass will help discourage the regrowth of surviving Poa trivialis and improve overall success of 

control (Morton and Reicher 2007).

4.3 Fish

Harvesting of invasive fish is generally only effective if the species is of importance to fisheries 

and anglers. Even when Bighead and Silver Carp were considered for harvest, harvest was found

to be one of the least effective methods available (Linfield 1980, Vacha 1998, Koehn et al. 2000, 

Wedekind et al. 2001, Arlinghaus and Mehner 2003).

Physical removal of fish can be an effective control technique in small ponds and other 

confined/constrained systems. Fish have been captured by drawdown, netting, gillnetting, 

trapping, and electroshocking. Goldfish and carp have frequently been managed in this way (Ritz 

1987, Morgan et al. 2005, Pinto et al. 2005). Fine-mesh monofilament gill nets have been used to 

control Rudd in three shallow lakes in Waikato, New Zealand, but elimination was not 

achievable (Neilson et al. 2004). Small, potentially fecund fish in dense littoral vegetation proved 
57

challenging to net, presenting a problem for total eradication, but removal of larger Rudd likely 

affected breeding success and netting is seen as a highly cost effective control method with low 

environmental impact (Neilson et al. 2004). Common Carp display jumping behavior when 

trying to escape entrapment. The Williams cage exploits this behavior by selectively removing 

the jumping carp from other fish (Stuart et al. 2011). Tests of the Williams cage in Australia 

proved to be extremely successful. Over a three-year test peroid, the Williams cage successfully 

separated 88% of adult Common Carp and allowed 99.9% of native species to pass through. The 

Williams cage is useful in controlling dispersal and abundance of Common Carp. The United 

States National Park Service uses physical removal through electrofishing to manage Rainbow 

Trout and Brown Trout populations that threaten native Brook Trout in Shenandoah National 

Park, Virginia (NPS 2011).

Barriers of several types have been used to control the spread of fish species and to prevent 

upstream migration or otherwise limit their access to spawning grounds. When using physical 

deterrents as barriers, combining methods can increase effectiveness. For example, Patrick et al. 

(1985) found that pelagic estuarine and freshwater fishes were successfully deterred by a barrier 

combining air bubbles and strobe lights.

Barriers and traps have been effective controls of Sea Lamprey since the 1950s. Barrier options 

include mechanical weirs, electrical barriers, low-head barriers, adjustable crest barriers, and 

velocity barriers (Scott and Crossman 1973, Smith and Tibbles 1980, GLFHC 2000ab). Traps are 

often used in association with barriers to capture Sea Lamprey while allowing desired species to 

continue upstream (GLFHC 2000a, FOC 2009). 

Barriers including electric, bubble curtain, and sonic have been used to exclude Common Carp 

from industrial cooling intake structures (Koehn et al. 2000). When possible, Common Carp can 

be excluded from an area and then kept out through sorting of fish (allowing desirable species to 

pass a barrier), which has been done since 1997 at the Cootes Paradise Marsh in Hamilton, 

Ontario (Lougheed et al. 2004). Although labor intensive, this method has proven effective at 

keeping Common Carp from returning to the marsh.

Electrical barriers and phonotaxis traps (cages with conspecific acoustic calls which allure gravid 

females) may be successful at limiting the movement of Round Goby. In tank studies, Round 

Gobies did not move through a barrier (Savino et al. 2001). Rollo et al. (2007) reported Round 

Gobies will approach a speaker emitting conspecific male calls in the field, and female round 

gobies showed significant attractions to speakers emitting conspecific male calls in the 

laboratory. Therefore phonotaxis could be used to lure gravid females to traps. As Round Gobies 

will spawn multiple times throughout late spring and summer (Jude et al. 1992), they should 

remain receptive to male calls and bioacoustic capture for the entire breeding season. 

Homing species such as salmonids typically return to their native stream for reproduction. 

Barriers can be constructed and/or natural barriers augmented to prevent upstream migration and 
58

aid management and eradication efforts. Lintermans and Raadik (2003) noted three key aspects 

of successful barriers to Rainbow Trout migration: there is a 1.5 m or greater vertical drop from 

the barrier to the pool below; in higher flows, the direction of water flow is towards the middle 

of the barrier with no slower overland flow passing down the banks; and no deep pool below the 

barrier from which fish could jump.

The United States Army Corps of Engineers Great Lakes and Mississippi River Interbasin Study 

(USACE GLMRIS) noted the potential effectiveness of sensory deterrent systems in providing 

barriers to fish migration or eliciting fish movements (USACE 2012b). Specifically, Maiolie et 

al. (2001) reported success using underwater strobe lights to deter wild, free-ranging 

Oncoryhnchus nerka in two large clear Idaho lakes an average distance of 30-136 m away, with 

an 80% reduction in fish density within 30 meters of the strobe lights. Models demonstrated 

strobe lights could be a successful deterrent to entrainment of Rainbow Smelt through Oahe 

Dam, Lake Oahe, South Dakota (Hamel et al. 2008). Patrick et al. (1985) found that combining 

strobe lights with air bubbles in a barrier would increase effectiveness of a barrier to Rainbow 

Smelt and other pelagic fishes.

The GLMRIS study also noted the potential effectiveness of acoustic fish deterrents in 

controlling or deterring Proterorhinus semilunaris populations. Acoustic deterrents include 

continuous wave and pulsed wave technology, which use sound/pressure waves to influence the 

behavior of aquatic organisms. Similarly, sensory deterrent systems such as acoustic air bubble 

curtains, electric barriers, and underwater strobe lights may prove useful in controlling 

populations in waterways and small bodies of water, but there are no studies of their effects on P. 

semilunaris at the present time (USACE 2012b). 

Water level manipulation has been used to to disrupt spawning of Common Carp (Summerfelt 

1999) and to exclude them from spawning habitat (Lougheed and Chow-Fraser 2001). 

Yamamoto et al. (2006) noted that physical drawdown of water levels has significant negative 

effects on cyprinid spawning abilities in Lake Biwa, Japan. Carassius spp. and Cyprinus carpio

eggs were notably reduced after collection when water levels were lowered by 30 cm, and as 

little as a 10 cm reduction significantly reduced available shallow, litter-accumulated spawning 

areas (Yamamoto et al. 2006).

Chemical piscicides are commonly used to control or eradicate invasive fishes, especially in 

rapid response scenarios. It should be noted that chemical treatment will often lead to non-target 

kills, and so all options for management of a species should be adequately studied before a 

decision is made to use piscicides or other chemicals. Potential effects on non-target plants and 

organisms, including macroinvertebrates and other fishes, should always be deliberately 

evaluated and analyzed. The effects of combinations of management chemicals and other 

toxicants, whether intentional or unintentional, should be understood prior to chemical treatment. 

Other non-selective alterations of water quality, such as reducing dissolved oxygen levels or 

altering pH, could also have a deleterious impact on native fish, invertebrates, and other fauna or 
59

flora, and their potential harmful effects should therefore be evaluated thoroughly (Finlayson et 

al. 2002).

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides. Antimycin and rotenone are non-selective, and toxicity to 

other fishes and aquatic invertebrates will vary (USACE 2013). TFM (3-trifluoromethyl-4-

nitrophenol) and niclosamide (2’,5-dichloro-4’-nitrosalicylanilide) are registered as lampricides, 

but show some promise for control of other invasive fishes as well (USACE 2013).

Rotenone is arguably the best known and most commonly used of the registered piscicides. The 

International Joint Commission (2011) recommends rotenone for control of White Perch and 

Round Goby in rapid response scenarios. Rotenone is also effective against Rainbow Trout, but 

at much higher concentrations than antimycin (Gilderhus 1972, Finlayson et al. 2002). 

Antimycin A is a registered piscicide in the United States that is documented as highly effective 

against scaled fishes. Antimycin is most effective in small streams, shallow ponds, and alpine 

lakes where there is ample mixing and an adequate contact time can be achieved (Gilderhus 

1972, Finlayson et al. 2002). Antimycin does not seem to repel fish like rotenone, and rapidly 

breaks down by hydrolysis in natural waters (Finlayson et al. 2002). Disadvantages of antimycin 

include increasing ineffectiveness in waters with higher pH (>8), streams with significant 

gradients (80-150 m elevation drop), and large lakes where good mixing and contact time cannot 

be established (Finlayson et al. 2002). In a study on removal of toxic chemicals from water using 

activated carbon, Dawson et al. (1976) found that granular activated carbon is saturated by 

rotenone. Antimycin was efficiently absorbed and did not saturate carbon at similar effective 

doses (Dawson et al. 1976). Antimycin-impregnated baits have been used to target Common 

Carp (Rach et al. 1994).

TFM has been used primarily to kill larval lampreys in their stream habitats (Smith and Tibbles 

1980). Beginning in the late 1950s, Sea Lampreys have been successfully controlled using TFM. 

The lampricide has reduced the population by over 90% of the 1961 peak (Scott and Crossman 

1973). However, continued use of TFM is required to keep populations under control (Scott and 

Crossman 1973, Becker 1983). TFM is sometimes harmful to other fish (e.g., Walleye) as well as 

to the larvae of nonparasitic native lamprey species (Becker 1983). 

Bayluscide (niclosamide) treatments in deltas are also a widely used and an effective control of 

Sea Lamprey larvae (NYSDEC 2012a). Exposure to niclosamide is known to be toxic to all fish 

species at 0.5 mg/L after a 48-h exposure (Clearwater et al. 2008). Niclosamide has been used 

for control of aquatic snails, Zebra Mussels, and oligochaetes, and is also toxic to many crayfish, 

frogs, clams, algae, and other amphibian and fish species (Clearwater et al. 2008).

Boogaard et al. (2003) found that the lampricides TFM and niclosamide demonstrate additive 

toxicity when combined. Evaluation of the effects of common piscicides on Ruffe revealed that 

the lampricide 3-trifluoromethyl-4-nitrophenol (TFM) has potential for selective control of the 
60

species (Boogaard et al. 1996). Boogaard et al. (2003) found that Brown Trout are among the 

least sensitive fishes to the lampricides. Ruffe was three to six times more sensitive to TFM than 

both Yellow Perch (Perca flavescens) and Brown Trout (Boogaard et al. 1996). A cost benefit 

analysis of a United States Ruffe control program supported TFM as a promising chemical 

control (Leigh 1998). However, Dawson et al. (1998) suggest that TFM may have more 

application for treating entire bodies of water rather than localized areas because it tended to 

repel Ruffe in preference tests, allowing them to move to untreated areas. Bottom-release 

formulations of bayluscide and antimycin showed promise for effectiveness in treating localized 

concentrations of Ruffe (Dawson et al. 1998).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five 

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

Changes in pH are known to affect fish behavior. Ikuta et al. (2003) documented the effects of 

low pH on Sockeye Salmon, noting that salmon would not swim upstream into areas of pH lower 

than 6.0. Acute exposure to low pH levels can directly kill fish by discharge of sodium and 

chloride ions from body fluid, and sub-lethal levels can affect reproduction. In the case of 

Sockeye Salmon, weak acidic conditions of <pH 6 were enough to depress the prespawning 

behavior of swimming upstream. Increases in pH through liming are listed as a potential control 

method for Goldfish (Clearwater et al. 2008).

Biological controls have also been commonly employed to control invasive fish species. 

Techniques range from the general – e.g., ‘top-down’ control to influence predator-prey balance 

– to exploitation of species-specific interactions. Because the trade-off between fish species as 

agents of biological control is not directly correlated with consumption, management decisions 

involving shifts between species should not take consumption solely into account (Stewart et al. 

1981).

Interactions among Sea Lamprey, salmonids (including several nonindigenous species), and the 

prey base (especially Alewife and Rainbow Smelt) have been key to biological controls targeting 

these species. Managing each of these species significantly impacts the others. Pacific salmon 

have significant environmental, socio-economic, and beneficial effects in the Great Lakes and 

therefore integrated management is essential. Pacific salmon prey heavily upon Alewife and 

Rainbow Smelt; Alewives remain a key food source and crucial to the survival of Pacific salmon. 

Over the past several decades, Pacific salmon populations have fluctuated with fluctuating 

Alewife populations. The management response to Great Lakes Alewife overabundance and 

recurring die-offs was to invest in Sea Lamprey (Petromyzon marinus) control and planting of 
61

hatchery-reared Pacific salmonids (Oncorhynchus spp.) to re-establish top open-water predators 

(Kocik and Jones 1999, Hansen and Holey 2002). Older and larger fish tend to be most heavily 

affected by piscivores, while smaller and younger fish remain abundant (Hewett and Stewart 

1989). Rainbow Smelt are also a major component of Pacific salmon diet. The presence or 

absence of this species significantly alters predator-prey relationships and competition between 

native species. Several species of non-native salmonids have been introduced to the Great Lakes, 

beginning in the 1960s, to control invasive Rainbow Smelt (USACE 2012b). Rainbow Smelt is 

heavily consumed by Atlantic Salmon (Salmo salar), Lake Trout (Salvelinus namaycush), Brook 

Trout (Salvelinus fontinalis), Coho Salmon (Oncorhynchus kisutch), Chinook Salmon 

(Oncorhynchus tshawytcha), Rainbow Trout (Oncorhynchus mykiss), Brown Trout (Salmo 

trutta), Splake (Brook Trout x Lake Trout), Burbot (Lota lota), Walleye (Sander vitreus), 

Northern Pike (Esox lucius), and many other freshwater piscivores (Stewart et al. 1981, Brandt 

and Madon 1986, Crossman 1991, He and LaBar 1994, Kirn and LaBar 1996, USACE 2012b). 

Observed Atlantic Salmon predation on smaller Rainbow Smelt, as well as bioenergetics 

modeling suggesting that by age 4, cumulative piscivory by Atlantic Salmon was nearly 10-fold 

greater than that of Lake Trout of the same age, implies its greater usefulness for management of 

Rainbow Smelt (Kirn and LaBar 1996). While Lake Trout consume large amounts of Rainbow 

Smelt, almost exclusively so in some studies, the species is believed to provide little potential for 

responsive management manipulation outside of stabilizing fluctuating prey populations, due to 

the long cycle of its predatory effect (peaking 3-5 years after stocking, lasting 7-8 years) (Stewart 

et al. 1981, He and LaBar 1994, Kirn and LaBar 1996). Chinook Salmon have been successfully 

used to eradicate Rainbow Smelt from small lakes in New Hampshire in 1936 (Stewart et al. 

1981). Of the 23 nonindigenous diseases and parasites in the Great Lakes, Aeromonas 

salmonicida, Renibacterium salmoninarum, Myxobolus cerebralis, and Novirhabdovirus sp. 

infections have been realized in Great Lakes Pacific salmon, while Heterosporosis sp. and 

Piscirickettsia cf. salmonis infections have been realized clinically or outside the Great Lakes. 

Glugea hertwigi, a microsporidian, is known to cause mortality in rainbow smelt. Therefore, 

Pacific salmon management must include the management of these pathogens and parasites.

Minnesota and Wisconsin, with advice from the U.S. Fish and Wildlife Service, implemented a 

top-down control program for Ruffe in the St. Louis River, western Lake Superior, in 1989, 

using Northern Pike (Esox lucius), Walleye (Sander vitreus), Smallmouth Bass (Micropterus 

dolomieui), Brown Bullhead (Ameiruus nebulosus), and Yellow Perch (Mayo et al. 1998). A 

bioenergetics modeling evaluation of the top-down control program revealed that although 

predators ate as much as 47% of Ruffe biomass in one year, they avoided Ruffe and were 

selective for native prey, and were thus unable to halt the increase in Ruffe abundance. However, 

the authors noted that Northern Pike and Walleye appeared to have potential for top-down 

control of ruffe due to a combination of their diets and population sizes, and due to indications 

that they may learn to prey more selectively on Ruffe. As Mayo et al. noted, caution is advised 

when considering top-down biological control as a management tool because the stability 
62

properties of a system do not just depend on predation, but also on the life histories of 

component species and their interactions.

Bottom-up control (reduction in food supply) of White Perch usually results in stunting 

accompanied by an increase in population so that the population consists of many small fish 

(Smith et al 2002).

Northern pike, Esox lucius, have been used as a biological tool to control Common Carp 

recruitment in the Sandhill lakes in Nebraska (Paukert et al. 2003). Great Lakes native Burbot 

were reported to depress abundance of Round Goby in eastern Lake Erie (Madenjian et al 2011).

An effective bio-control of Sea Lamprey is the implementation of the sterile-male-release 

program. Male Sea Lamprey are captured during spawning runs, sterilized using bisazir, and 

released to compete with fertile males for mating; thus reducing egg fertilization. Released males 

are sterilized past their parasitic phase and do not return to the lake. (FOC 2009, GLFHC 2000a). 

Application of different pheromones such as migratory, alarm, and sex may be useful in the 

integrated management of carp (Sorensen & Stacey 2004) as well as Sea Lamprey.

Inducible Fatality Genes (IFG) involve breeding fishes with a fatal genetic weakness to a trigger 

substance, such as zinc. The fatal gene technology appears to be a potentially viable and longterm strategy for the environmentally benign control of carp (Koehn et al 2000).

4.4 Mollusks

Eradication of mollusks from infested open waters is unlikely – emphasis is generally on 

preventing further spread, limiting population impacts (especially relating to infrastructure such 

as water intakes), and controlling new infestations in small water bodies (e.g., ponds, water 

gardens).

Physical controls can be an important component in the control of invasive mollusks. With the 

possible exception of the large ‘escargot’ snail species (Cipangopaludina and Viviparus spp.), 

there is not a significant harvest of any of the Great Lakes nonindigenous mollusk species and 

harvest is not consider to impact control. Physical removal has little effect on control in 

ecosystems, but can be employed to protect infrastructure (e.g., water intakes, swimming 

beaches), usually in conjunction with physical barriers. Screens and traps are commonly 

employed to prevent Corbicula colonization of water intakes (GISD 2013). Diver assisted 

suction removal and bottom barriers are being researched as potential methods for physical 

control of Corbicula populations in Lake Tahoe (UC Davis TERC 2004). Benthic barriers have 

been effective for short-term control of Corbicula fluminea, but non-target mortality to other 

benthic invertebrates may be high (Wittmann et al 2012). Effective physical controls of 

Dreissena include infiltration intakes or sandfilter intakes (to filter out veligers), thermal 

treatments, carbon dioxide pellet blasting, high-pressure water jet cleaning, mechanical cleaning, 

freezing, scraping, scrubbing, pigging, and desiccation. Placement of water intakes in areas too 

deep or otherwise unsuitable for zebra mussel colonization has been used as a form of physical 
63

control, but this is less successful in areas that also have Quagga Mussels. Other potential 

dreissenid controls include the use of electrical fields, pulse acoustics, low-frequency 

electromagnetism, ultraviolet light (UV light), and reduced pressure (USACE 2002). New 

Zealand Mudsnail control in hatcheries has included using a flame thrower against the walls of 

raceways, since mudsnails cannot withstand warm temperatures (Dwyer et al. 2003, Richards et 

al. 2004) or low humidity situations (Dwyer and Kerans, unpublished, Richards et al. 2004). 

New Zealand Mudsnails are also sensitive to freezing and can be eradicated if an infested area is 

drained in the winter, and the substrate is frozen to the depth containing the mudsnails. There is 

preliminary evidence that hydrocyclonic separators may be a useful tool to decontaminate fish 

hatchery water supplies and prevent the spread of New Zealand Mudsnails within a hatchery.

Physical removal of visible vegetation (which may harbor invasive mollusks) from boats, 

trailers, and other equipment being moved from one water body to another is an important 

method in controlling spread. Asian clam, Corbicula fluminea, may be controlled at intake pipes 

by heating influent water to 99oF (GISD 2013). Flushing engines, cooling systems, live wells,

and bilge with water over 110oF will kill dreissenid veligers and 140oF will kill adults. The New 

Zealand Mudsnails can survive at 43.3°C, so the water temperature needs to exceed 120oF to 

eliminate that species (Medhurst 2003). Drying fishing gear at 28.9-30°C for at least 24 hours or 

at 40°C for at least two hours will eliminate New Zealand Mudsnails (Richards et al. 2004). Air 

drying equipment for 5 days will kill most dreissenid larvae and smaller dreissenid mussels, but 

large mussels may survive up to two weeks out of water. Dessication (drying) is not an effective 

control method for Cipangopaludina chinensis. Field experiments under mesic conditions 

indicated that this snail can survive exposure to air for at least 4 weeks (Havel 2011). Putting 

fishing gear in a freezer for 6-8 hours will kill all attached NZ mudsnails (Medhurst 2003, 

Richards 2004). 

Preliminary research demonstrates that Cipangopaludina chinenesis will not migrate upstream 

against a small current (Rivera 2008). Authors suggest that acceleration of current may be an 

important management tool for preventing upstream spread.

Many chemical molluscicides are available, but are not species-specific and may harm native 

species to a greater extent than non-natives. Many of the nonindigenous mollusk species can 

close their shells tightly, reducing the effectiveness of chemical controls. In short-term 

experiments, Sphaerium corneum can reduce the bioaccumulation of 2,4,5-trichlorophenol (TCP) 

by closing their shell valves (Heinonen et al 1997) – this reaction to chemical stimuli generally 

may limit the usefulness of chemical molluscicides against this species. Likewise, 

Cipangopaludina spp. can close their operculum tightly and have been shown to be more 

resistant to chemical treatment than native snails without that ability.

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N,N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 
64

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012).

Chlorination has been the most common treatment for control of Dreissena mussels, but chlorine 

concentrations needed for effective control of quagga mussels may reach hazardous levels 

(Grime 1995). Oxidizing chemical control treatments effective against D. polymorpha include 

hypochlorite, chloramine, chlorine dioxide, bromine, ozone, potassium permanganate, and 

sodium chlorite. 

Corbicula is not tolerant of fluctuating environmental conditions (particularly temperature and 

oxygen) and is prone to massive die-offs (Menninger 2013), this suggests that short-term 

chemical manipulation may be useful in controlling Corbicula populations. Low concentration of 

chlorine or bromine will kill juvenile Asian Clams (GISD 2013).

Chemical methods used to eradicate New Zealand Mudsnails include: Bayer 73, copper sulfate, 

and 4-nitro-3-trifluoromethylphenol sodium salt (TFM). The only molluscicide known to have 

been tested against New Zealand Mudsnails is Bayluscide (niclosamide). Hyamine and hydrogen 

peroxide have also been used to control New Zealand mud snails (IJC 2011). Preliminary 

investigations also suggest that copper and carbon dioxide under pressure may prove useful in 

both decontaminating fish hatchery water supplies and preventing spread into uncontaminated 

areas of a hatchery. Ozone has not been shown to be effective in killing New Zealand Mudsnails 

in a hatchery environment.

Various chemical coatings – including copper-based, tributyltin-based, copolymer, vinyl/epoxy, 

resin and other films - can be applied to structures to prevent the attachment of Zebra Mussels. 

Tributyltin-based antifoulants are extremely toxic and restricted by federal law (Ohio Sea Grant 

1992).

Biological control so far has proven to be ineffective in controlling Dreissena species. Predation 

by migrating diving ducks, fish species, and crayfish may reduce mussel abundance, though the 

effects are short-lived (Bially and MacIsaac 2000). Other biological controls (biopesticides)

being researched are selectively toxic microbes and parasites that may play a role in management 

of Dreissena populations (Molloy 1998). Laboratory testing shows strain CL145A of 

Pseudomonas fluorescens (a bacterium) to be highly lethal to Zebra Mussels; capable of 

eliminating over 90% of adults and 100% of larvae (Molloy 2002, Abdel-Fattah 2011). 

Commercially, this product is known as Zequanox® and is developed by Marrone Bio 

Innovations (Abdel-Fattah 2011) – this product has been registered by EPA for use as a 

pesticide. Interfering with the synchronization of spawning by adults in their release of gametes 

could also offer control of Dreissena populations (Snyder et al. 1997). Another approach would 

be to inhibit the planktonic veliger (larvae) from settling and attaching to a surface to begin 

development (Kennedy 2002).

Competition with dreissenid mussels will likely limit expansion of Lasmigona subviridis and 

other large nonindigenous bivalves in the Great Lakes.
65

Manipulation of predator fishes and turtles that eat snails may be useful in the control of snail 

populations. However, as a relatively large snail species, Cipangopaludina chinensis, may 

escape predation by smaller fishes. Parasites of New Zealand Mudsnails from New Zealand may 

also become useful to control population size by inhibiting reproduction. Studies of the efficacy 

and specificity of a trematode parasite from the native range of New Zealand Mudsnails as a 

biological control agent have shown positive results so far (Dybdahl et al. 2005).

4.5 Insects

Acentria ephemerella is used for biological control of Eurasian Watermilfoil (Myriophyllum 

spicatum). Its population is best controlled by elimination of its host plants – which are 

predominantly Eurasian Watermilfoil but also may, to a lesser extent, include a variety of other 

native and nonindigenous plants (Cornell 2004). Tanysphyrus lemnae is an herbivore of 

duckweeds and other closely-related aquatic plants. It is likely also best controlled through 

control of its host plants. 

4.6 Free-Living Crustaceans, including Waterfleas, Copepods, Mysids, and Gammarids

Microcrustaceans are easily transported overland by recreational boaters and on fishing gear. 

Species with large tail spines, such as Bythotrephes and Cercopagis are especially likely to be 

spread in this way, as are resting eggs that are adapted to promote attachment (as in the hooks on 

Daphnia lumholtzi epphipia). Fishing lines designed specifically to prevent the spread of 

waterfleas, such as the Flea Flicker brand, have been proven effective in significantly reducing 

fouling on lines, indicating their importance as a management tool (Jacobs and MacIsaac 2007). 

Cleaning all aquatic equipment with high-pressure water (>250 psi) or hot water (>50°C) after 

each use has proven effective for controlling spread via this pathway. Echinogammarus ischnus

can tolerate a maximum water temperature between 31.0°C and 32.2°C and Gammarus tigrinus

between 32.2°C and 34.2°C before irreversible physiological damage and mortality occur 

(Wijnhoven et al. 2003). Hemimysis anomala also exhibits mortality at temperatures below 0° C 

(Borcherding et al. 2006), making freezing of gear an alternative.

Electron beam irradiation has been used to control microorganisms in aquatic pathways, 

including invasive waterfleas and copepods (USACE 2012b). Electron beam irradiation is a nonselective control method which exposes water to low doses of radiation using gamma-sterilizers 

or electron accelerators, breaking down DNA in living organisms while leaving behind no byproducts (USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms in 

water treatment facilities and narrow channels, where UV filters can be used to emit UV light 

into passing water, penetrating cell walls and rearranging DNA of microorganisms (USACE 

2012b). Many ballast water treatment systems already available and/or under development use 

UV as a primary biocide, usually preceded by filtration (Lloyd’s Register Marine 2014). Another 

possible non-selective pathway control is high water turbidity, which may decrease zooplankton 

(especially cladoceran) abundances due to the negative effects of suspended clay particles on 

filtering and assimilation rates (Suchy and Hann 2007).
66

Waterfleas are consumed by planktivorous fishes and predatory invertebrates; manipulation of

predator populations (top-down control) has been used to control waterflea populations. Youngof-year Bloaters (Coregonus hoyi) have been shown to surface feed on Eubosmina coregoni in 

Lake Michigan, but their effectiveness as a control is unknown, especially because they may

move to the benthos earlier to avoid competition from the invasive Alewife (Alosa 

pseudoharengus) (Crowder and Crawford 1984). Bythotrephes spp. are known to consume E. 

coregoni in Russia and the United States (Grigorovich et al. 1998), and E. coregoni populations 

declined significantly immediately following the invasion of the waterflea Bythotrephes 

longimanus in the Laurentian Great Lakes Michigan, Huron, and Erie, with direct predation by 

B. longimanus the most likely explanation (Barbiero and Tuchman 2004). E. coregoni levels in 

B. longimanus-invaded areas have remained at low levels, indicating a significant populationwide impact, but the long-term effectiveness of B. longimanus as a biological control method is 

unknown (Barbiero and Tuchman 2004). B. longimanus is consumed by Rainbow Smelt, Lake 

Herring, Lake Whitefish, Yellow Perch, White Perch, White Bass, Walleye, Alewife, Bloater 

Chub, Emerald Shiner, Spottail Shiner, Deepwater Sculpin, and Chinook Salmon in the Great 

Lakes (Bur et al. 1986, Makarewicz and Jones 1990, Branstrator and Lehman 1996). The

defensive tailspine on Bythotrephes longimanus has been observed to increase in size throughout 

the summer in response to predation pressure (Straile and Halbich 2000). Consequently, larger 

fish are more likely to be successful predators (Branstrator and Lehman 1996). The opossum 

shrimp (Mysis relicta) has been observed eating B. longimanus in Ontario lakes, but the 

frequency of consumption appeared related to abundance of the invader and alternate prey 

(Nordin et al. 2008). Pothoven et al. (2007) found that adult large Alewives (Alosa 

pseudoharengus) (>100 mm) consume Cercopagis pengoi in Lake Michigan, but not 

significantly enough to control the species, concluding that the Alewife prefers Bythotrephes 

longimanus due to its larger size and conspicuousness. In contrast, a study of C. pengoi as a prey 

item in Lake Ontario found that at least 70% of Alewives larger than 70 mm contained C. pengoi

spines (Bushnoe et al. 2003). The same study also found spines in Rainbow Smelt (Osmerus 

mordax) stomachs. Rainbow Smelt historically consume cladocerans in the Great Lakes, but 

prefer larger prey and may select B. longimanus over C. pengoi where both occur (Pothoven et 

al. 2009). Gorokhova et al. (2004) found that in the northern Baltic proper, Herring (Clupea 

harengus L.) and Sprat (Sprattus sprattus L.) are the dominant predators of C. pengoi, and a 

possible source of biological control through fisheries management, though it is possible that 

fully mature resting eggs may survive passage through fish digestive systems as has been 

observed with B. longimanus eggs in Yellow Perch (Perca flavescens). B. longimanus is also 

known to consume C. pengoi, but not as a main prey item (Cavaletto et al. 2010). Daphnia 

lumholtzi is preyed upon by a variety of zooplanktivorous fishes, including Inland Silversides, 

Menidia beryllina, in Lake Texoma, Bluegill (Lepomis macrochirus), White Bass (Morone 

chrysops), White Crappie (Pomoxis annularis), and Black Crappie (Pomoxis nigromaculatus) in 

Lake Chautauqua, Illinois (Lienesch and Gophen 2001, Lemke et al. 2003). The degree to which 

these fishes may be able to control Daphnia lumholtzi populations is not certain. The mysid 
67

shrimps Mysis mixta and Mysis relicta consume Eubosmina maritima diapausing eggs (ephippia) 

selectively in the northern Baltic Sea, but only at a rate which can affect local abundances of E. 

maritima (Viitasalo and Viitasalo 2004). Research on D. galeata galeata is lacking, but many 

invertebrates are likely predators of Daphnia spp. where they occur in Europe and North 

America, including Great Lakes species such as the predacious phantom midge Chaoborus 

flavicans, the waterfleas Leptodora spp. and B. longimanus, (Lysebo 1995). Many small fish of 

species such as Yellow Perch (Perca flavescens), Three-spined Stickleback (Gasterosteus 

aculeatus), Alewife, Bluegill, and Ciscos have been documented consuming daphnids and other 

zooplankton in Canada and the United States (Mills and Forney 1983, Post and McQueen 1987, 

Hulsmann and Mehner 1997).

Free-living copepods are also preyed upon by fish and invertebrate predators and top-down 

control may be a useful tool. Treasurer (1992) found that Eurasian Perch, Perca fluviatilis, larvae 

selectively prey on Cyclops strenuus abyssorum in the Scottish lochs Kinord and Davan. The 

total zooplankton reduction observed was minimal, but Treasurer suggested that grazing by 

larvae is likely to impact copepod populations. Maes et al. (2005) found that juvenile Herring 

(Clupea harengus) and Sprat (Sprattus sprattus) exhibit top down control of Eurytemora affinis

through predation pressure in the Scheldt estuary in Belgium. Yellow Perch (Perca flavescens) 

and Trout Perch (Percopsis omiscomaycus) are potential predators of Megacyclops viridis (Ogle 

et al. 1995). Hansen and Jeppesen (1992) found that a 50% reduction of planktivorous fish 

biomass (Roach, Rutilus rutilus, and Bream, Abramis brama) affected cyclopoid copepod 

population directly through reduction in fish predation pressure and changes in biological 

structure of Lake Væng, Denmark. Megacyclops viridis is also an important prey item for 

introduced Ruffe (Gymnocephalus cernuus), but the Ruffe feeds on a wide variety of benthic 

organisms, so its feasibility as a biocontrol is unknown (Ogle et al. 1995). Nitokra hibernica has 

been found in the stomach of Slimy Sculpin (Cottus cognatus) at Yankee Reef, Lake Huron, and 

it is known to be consumed by Rainbow Smelt (Osmerus mordax) in Lake Huron, but the 

significance of this predation on biological control is unknown (Lesko et al. 2003).

Benthic invertebrates including Echinogammarus ischnus are a major part of the native Yellow 

Perch (Perca flavescens) diet (Gonzalez and Burkart 2004). Echinogammarus ischnus has also 

become prey of the invasive Round Goby, Neogobius melanostomus (Gonzalez and Burkart 

2004). The spread of dreissenid-covered substrate across the Great Lakes region has created an 

ideal habitat for Echinogammarus ischnus, where it is less susceptible to predation, indicating 

that efforts to control Dreissena spp. could also aid control of Echinogammarus ischnus 

(Gonzalez and Burkart 2004). A parasitic water mold (oomycete) detected in the upper St. 

Lawrence River is likely responsible for reducing Echinogammarus ischnus abundance despite 

favorable physical and chemical conditions (Kestrup et al. 2011). The oomycete also infects the 

native amphipod Gammarus fasciatus, but its effects are significantly less severe than in 

Echinogammarus ischnus (Kestrup et al. 2011). In a study of the effects of tide gate operation on 

hypoxic conditions in the Back River and Savannah River estuaries in Savannah, Georgia, it was 
68

found that oxygen saturation levels below 30% are lethal to experimental populations of 

amphipods (Winn and Knott 1992). Electron beam irradiation can be used to control 

microorganisms in aquatic pathways, including Echinogammarus ischnus (USACE 2012b). 

Electron beam irradiation is a non-selective control method which exposes water to low doses of 

radiation using gamma-sterilizers or electron accelerators, breaking down DNA in living 

organisms while leaving behind no by-products (USACE 2012b). Ultraviolet (UV) light can also 

effectively control microorganisms including E. ischnus in water treatment facilities and narrow 

channels, where UV filters can be used to emit UV light into passing water, penetrating cell 

walls and rearranging DNA of microorganisms (USACE 2012b).

Various parasites have been shown to reduce host density and population survival in 

experimental Daphnia populations in Europe (Ebert 2005). However, it is unclear whether 

parasites regulate natural Daphnia populations, as all experiments have been completed under 

lab conditions. 

Zooplankton biomass generally increases with increasing eutrophication, so reduction of 

excessive nutrient pollution causing abnormal eutrophication could help control invasive 

waterfleas. Gemza (1995) documented a shift from copepods to cladocerans as dominant 

zooplankton at increasingly eutrophic sites in Severn Sound, Lake Huron. Many of the nonnative freshwater cladocerans that have invaded the Great Lakes are susceptible to salinities >3% 

(Nauwerck 1991). Santagata et al. (2008) found that ballast water exchange methods which flush 

freshwater organisms into euhaline seawater are effective against Eubosmina maritima at a 

minimum of 24 PSU (practical salinity units) for two hours in a laboratory simulation. However, 

E. maritima ephippia may remain in residual unpumpable sediment in ballast tanks, and Gray et 

al. (2005) found that exposing zooplankton ephippia to open ocean saline water of 32 ppt (parts 

per thousand) did not reduce egg abundances or consistently affect richness of invertebrates 

hatched from exposed eggs. Hemmimysis anomala tolerates salinity of 0-19 ppt (parts per 

thousand) (Bij de Vaate et al. 2002). Ellis and MacIsaac (2009) documented 100% mortality for 

H. anomala after five hours in a simultaneous BWE treatment, in which salinity was gradually 

increased from 4-30 ppt, and 100% mortality after three hours in a sequential BWE treatment, in 

which species are immediately exposed to 30 ppt salinity.

A study of the effects of cadmium and zinc on Lake Michigan zooplankton found that E. 

coregoni was significantly reduced by separate and combined treatments, with negative effects 

primarily due to zinc (Marshall et al. 1981). A more recent study on the effects of copper sulfate 

(used to control algal biomass in eutrophic water bodies) and Carbaryl (used to control aquatic 

pests) on zooplankton found that levels of 50 μg/L Cu and 20 μg/L Carbaryl individually 

reduced E. coregoni biomass by >50% (Havens 1994).

Lindley et al. (1999) found that exposure to the organochlorine compounds pentachlorophenol 

(PCP) and 1, 2-dichlorobenzene (DCB), (both common industrial pollutants) accumulated in 

sediment significantly reduced hatching success and nauplii viability of E. affinis eggs. In a study 
69

of the effect of salinity on toxicity of cadmium to Chesapeake Bay organisms, Hall et al. (1995) 

found that E. affinis is very sensitive to cadmium compared to other estuarine aquatic biota. The 

negative effects of the insecticide diflubenzuron on E. affinis nauplii were documented by Savitz 

and Wright. The insecticide is approved for use against the Gypsy Moth (Lymantria dispar) and 

other insect pests by the United States Environmental Protection Agency, and enters E. affinis

habitat primarily through runoff (Savitz and Wright 1994). Diflubenzuron specifically targets the 

arthropod molting process, so the most explicit effects are expected in sub-adult crustaceans.

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of crustaceans.

4.7 Free-Living Worms including Annelids and Playthelminthes

Little research has been pursued concerning control of aquatic annelids. The effects of industrial 

toxicants on Tubificidae species and explorations of their value as an indicator of environmental 

quality have been explored, but chemicals and heavy metals are not viable methods of control 

because of unknown and adverse effects on the surrounding environment (Das and Das 2005, 

Saha et al. 2006). However, there has been investigation into the control of Branchiura sowerbyi

as a host of haemorrhagic thelohanellosis, which negatively impacts fish in aquaculture 

(Liyanage et al. 2003).

Brown Trout, Salmo trutta L., has been shown to prey on oligochaetes; its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, Brown Trout is itself an invasive species in the Great Lakes region and across nearly 

all of the United States.

Research on benthic macroinvertebrate communities in southwestern Lake Ontario before and 

after the invasion of Dreissena polymorpha (Zebra Mussels) and Dreissena bugensis (Quagga 

Mussels) suggests that the presence of Dreissena helps to improve benthic habitat, facilitating 

increases in macroinvertebrates, including the tubificids Potamothrix vejdovskyi and 

Spirosperma ferox (Stewart and Haynes 1994). This indicates that control of invasive Quagga 

and Zebra Mussels could facilitate improved control of benthic macroinvertebrates such as the 

tubificids.

Researchers found that Branchiura sowerbyi, which is a vector in transmission of Thelohanellus 

hovorkai (myxozoa) to fish, prefers muddy substrate, while other benthic oligochaetes that are 

not susceptible to myxozoa prefer sandy substrate, and suggested that replacing bottom substrate 

from mud to sand would lead to a shift in oligochaete communities from Branchiura sowerbyi to 

non-susceptible oligochaetes such as Limnodrilus socialus, therefore reducing disease in 

aquaculture fauna (Liyanage et al. 2003).
70

Potamothrix bedoti has been shown to be more likely to occur in substrate with a high clay and 

silt content, (Sauter and Gude 1996) while Potamothrix moldaviensis has been shown to be more 

likely to occur in sandy substrate with a clay and silt content of less than 10%. This indicates that 

substrate type is a possible physical control method to be further explored.

Ripistes parasita has been found to occur in greater numbers where water quality is impaired by 

industrial pollution, therefore greater measures to control pollutants such as heavy metals and 

particulate matter might help control this oligochaete (Simpson and Abele 1984). Furthermore, 

declines in Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in 

correlation with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of 

excess nutrients would help to reduce oligochaete populations.

Only one free-living nonindigenous platyhelminth is found in the Great Lakes, Dugesia 

polychroa. No research is available supporting control of this species.

4.8 Bryozoans, Hydrozoans, and Testate Amoebae

Lophopodella carteri colonies grow on solid substrata (Lauer et al. 1999), therefore, physical 

removal methods such as scraping may be viable for localized areas. Chemical biocides have 

been used as anti-fouling agents to remove sessile macroinvertebrates from shipping equipment 

and industrial intakes, but none have been approved for use on bryozoans as of yet (USACE 

2012b). The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that 

alteration of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could

be effective in preventing upstream and downstream movement of bryozoans. Freshwater 

Bryozoans are generally sensitive to heavy metals, particularly copper (Bushnell 1974). Pardue 

and Wood (1980) determined baseline toxicity of four heavy metals to three species of 

phylactolaemate bryozoa (L. carteri, Pectinatella magnifica Leidy, and Plumatella emarginata

Oka), observing greatest sensitivity to cadmium, followed by copper, chromium, and zinc. It 

should be noted that the toxicity of these metals were not tested as control measures, but as a 

demonstration of the usefulness of some bryozoans as biomonitors of water quality. However, 

comparison of the 96-hr LC50 data to toxicity data from other studies indicates that the bryozoa 

are more sensitive to the tested metals than many other invertebrates and fish, indicating 

potential as chemical controls with further research (Pardue and Wood 1980). It should be noted 

that at least one invasive Marine Bryozoan (Bugula neritina) has demonstrated heavy metalresistant genotypes, suggesting that metal-intensive antifouling agents may have diminished 

effectiveness on their populations (Piola and Johnston 2006).

Control of Cordylophora caspia will likely focus on its potential role as a biofouling agent. 

Cordylophora spp. has been documented colonizing the inner walls of power plants in Europe 

and the United States (Folino 2000), primarily causing blockages in nozzles and tailpipes of 

rapid gravity filter beds (RGFs) (Mant et al. 2011). The menont life-stage of C. caspia often 

found in hydroelectric intakes, is both drought and temperature resistant which may prove an 
71

obstacle to control (Gutierre and Gutierre 2012). Thermal treatments greater than 37°C have 

been proven effective in eradicating colonies of Cordylophora spp. sampled from the walls of 

power plant intakes (Folino 2000), but are not efficient in water treatment facilities where there 

is no residual heat energy available (Mant et al. 2011). Gutierre and Gutierre (2012) found that 

Cordylophora caspia is completely eradicated at pH levels of 4.0 and 10.0, with increasing 

survival rates in between, and suggested maintaining pH levels at 10.0 for six hours or more by 

injection of NaOH to reduce and eliminate colonies. Chlorine treatments negatively affect 

Cordylophora growth, but treatments as high as 5 mg/L for periods of 105 minutes have been 

unsuccessful in completely eradicating colonies (Mant et al. 2011). Furthermore, chlorine use is 

highly regulated at water treatment facilities where Cordylophora most frequently cause 

problems. Hydroids are sensitive to vanadium leeching from slag stones used in riverbank 

reinforcement, and are sensitive to heavy metals in general, especially mercury, copper, 

cadmium, and arsenic, though it is unlikely that these will be useful in control (Ringelband and 

Karbe 1996).

Craspedacusta sowerbyi has spread across temperate climates for more than a century, but 

despite experimental observation of its possible contribution to trophic cascade effects 

(Jankowski et al. 2005), and studies on predation habits (Dendy 1978, Dodson and Cooper 1983, 

Spadinger and Maier 1999), little research on control is available.

Three nonindigenous members of Genus Psammonobiotus have been doucumented as 

established in the Great Lakes. No research is available supporting control of these species.

4.9 Parasites and Diseases

Most control research conducted for nonindigenous fish parasite and disease species has been in 

the context of aquaculture and recreational fisheries pond management. Minimizing fish stress 

can reduce the risk of disease outbreak (FTS 2012). No treatments exist to control these diseases 

and parasites in open systems. Establishment of quarantines may prevent transmission and be 

useful in controlling spread.

4.9.1 Parasitic Platyhelminthes

Implementation of Eurasian Ruffe management may potentially decrease Dactylogyrus spp.

prevalence due to host specificity. Lampricide TFM may effectively eliminate up to 97% of 

Ruffe, potential carriers of Dactylogyrus amphibothrium, with minimal non-target mortality 

(Crosier et al. 2012). However, Ruffe management is considered by some (e.g., Ogle 1998) to be 

difficult and impractical given that the species has developed several adaptations to compensate 

for high mortality rates (Lind 1977) and populations rebound quickly (Lelek 1987). 

Bothriocephalus acheilognathi populations in aquaculture and ponds can be controlled by 

managing the intermediate host (i.e., copepods) population densities. Effective ectoparasiticides 

include Neguvon®, Masoten®, Dipterex®, Bromex®, and Naled® (Paperna 1996).
72

Bath treatments are effective control methods for Bothriocephalus acheilognathi infections. 

Baths should contain Droncit® (praziquantel), isopropyl alcohol, and water yielding a final 

mixture concentration ≥ 0.67 ppm praziquantel. Fish densities during treatment should be no 

greater than 60 mg fish/L and exposure should last 24 hours. After 24 hours, the treatment should 

be drained, worm parts discarded, and clean water added. Fish should then be transferred to a 

decontaminated container (Mitchell and Darwish 2009).

Dactylogyrus-specific treatments are unknown. However, multiple chemicals are effective at 

treating monogenean fluke infections in aquaculture systems. Effective benzimidazoles include 

levamisole (Buchmann 1997) and praziquantel, which has high efficacy against Dactylogyrus

spp. (Schmahl and Mehlhorn 1985, Buchmann 1997). Effective bath treatments include 

formaldehyde (30-100 ppm), sodium chloride, copper sulphate, hydrogen peroxide, sodium 

percarbonate (Buchmann and Kristensen 2003), formalin (25 mg/L for prolonged exposure or 

150-250 mg/L for 30 minutes), and potassium permanganate (2 mg/L for prolonged exposure or 

10 mg/L for 30 minutes) (Reed et al. 1996). Effective organophsophate bath treatments include 

metrifonate (0.25-0.5 ppm) and dichlorvos (0.25-0.5 ppm) (Sarig et al. 1965). Pond infestations 

can be controlled with formalin (30 mg/L) or trichlorfon (Lepidex®; 0.5 mg/L) (Reed et al. 

1996). However, monogenean eggs display chemical resilience and therefore the above chemical 

treatments are ineffective at destroying eggs (Reed et al. 1996, Rowland et al. 2007). Chemical 

toxicity varies considerably between monogeneans and fish species. Toxicology and tolerance 

tests are suggested prior to using anthelmintics (“dewormers”). Managers are encouraged to 

consider specific host drug tolerance, temperature, salinity, organic material content, and drug 

retention time prior to treatment (Buchmann and Bresciani 2006). Freshwater fish species can 

also be dipped in saltwater to minimize external parasite numbers prior to stocking (Reed et al. 

1996).

Bothriocephalus acheilognathi infections can be treated with chemically enhanced feed. Drugs 

should be mixed in oil and sprayed on feed at a rate of 1 L/70 kg dry weight. Effective chemicals 

and doses include dibutylin oxide or dibutylin dilurate (250 mg/kg fish) fed over three days 

(Mitchell and Hoffman 1980), Yomesan® (500 g/500 kg dry pellets) fed at 1.5% of body weight 

2–3 times weekly, and Yomesan® (28 g/40 kg) fed for three days (Korting 1974, Mitchell and 

Hoffman 1980, Brandt et al. 1981).

No effective treatments have been identified for Ichthyocotylurus pileatus, Neascus 

brevicaudatus, Scolex pleuronectis, or Timoniella sp.

4.9.2 Parasitic copepods

Electron beam irradiation can be used to control microorganisms in aquatic pathways, including 

Neoergasilus japonicus (USACE 2012b). Electron beam irradiation is a non-selective control 

method which exposes water to low doses of radiation using gamma-sterilizers or electron 

accelerators, breaking down DNA in living organisms while leaving behind no by-products 
73

(USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms including N. 

japonicus in water treatment facilities and narrow channels, where UV filters can be used to emit 

UV light into passing water, penetrating cell walls and rearranging DNA of microorganisms 

(USACE 2012b).

Gault et al. (2002) documented the effectiveness of “egg-laying boards” in control of the 

confamilial Argulus foliacus in a Rainbow Trout (Oncorhynchus mykiss) fishery. Lightweight, 

corrugated polypropylene boards were anchored and submerged within six mm of the water 

surface in a horizontal plane or floated vertically at varying depths to provide ideal eggdepositing surfaces for A. foliacus. Replacement of the boards at two week intervals and constant 

deployment throughout the A. foliacus breeding season significantly reduced the prevalence and 

intensity of parasite infection in trout, but the boards do not stop all egg hatching due to clutches 

laid on other pond surfaces. The most effective method of removing Argulus spp. from 

recreational fisheries and aquaculture ponds is complete drainage and removal/restocking of fish, 

possibly combined with a treatment of lime, but this is not economically or physically possible in 

many cases (Gault et al. 2002, Taylor et al. 2006).

The organophosphate compound Dipterex (organophosphate trichlorphon) has been used to 

control argulids in ponds and aquaculture (Tonguthai 1997). Argulid adults and larvae fall off 

parasitized fish and die within 12-24 hours after treatments of 0.2-0.3 ppm, sprinkled or sprayed 

uniformly on the pond surface (Tonguthai 1997). However, argulid resting stages are not 

affected by Dipterex—multiple treatments must be applied over the hatching period for effective 

reduction. In a study on chemotherapeutic control of ectoparasites in aquaculture, Singhal et al. 

(1986) found gammexane (1-6-hexachlorocyclohexane) to be most effective in removing 

Argulus indicus from host fish Indian Carp (Catla catla), Rohu (Labeo rohita), Mrigal Carp 

(Cirrhinus mrigala), Grass Carp (Ctenopharyngoden idella), and Silver Carp 

(Hypophthalmichthys molitrix) after a treatment was sprayed over the pond surface at 0.2 mg L-1 

three times at weekly intervals. Placing Catla catla, L. rohita, and Ctenopharyngoden idella in 

30 mg L-1 sodium chloride solution for 2-5 minutes was effective in removing A. indicus, and 1-

2 minutes sufficed for H. molitrix and Cirrhinus mrigala (Singhal et al. 1986). Placement of fish 

in 0.5 mg L-1 potassium permanganate achieved 60% reduction of parasites within five minutes. 

For severe infestations, 0.2 g L-1 of lime was used to treat drained, dried ponds, after which they 

were restocked. Combinations of organophosphate pesticides and salt are reported to have 

controlled Argulus japonicus after four treatments at one week intervals (Avenant-Oldewage 

2001). As with Dipterex, it should be assumed that the above treatments are ineffective against 

argulid resting stages, as the effects have only been studied on hatched copepods.

4.9.3 Protozoan Parasites

Effective physical control methods for Heterosporis include complete desiccation of holding 

tanks and equipment for 24 hours, freezing at -20°C for 24 hours, and culling (Sutherland et al. 

2006, GLFHC 2012b). Immersion of gear in a 2200 ppm bleach (0.7 L bleach per 20 L of water) 
74

solution for five minutes will destroy the parasite (IDNR 2005, Sutherland et al. 2006, GLFHC 

2012b).

Exposing myxospores (Myxobolus cerebralis) to 90°C water for 10 minutes is effective at 

destroying the spores (Hoffman and Markiw 1977). Electrical charges of 1-3 kV pulsed 1-25 

times at 99 μsec/pulse is effective at killing large numbers of TAM spores (Wagner 2002). 

Experiments by Hoffman (1974) have demonstrated that filtration is not an effective method for 

removing TAM spores from water – due to the small spores size, the filter needed to remove 

them slows flow to rates unacceptable for most applications. Hatchery intake water treated with 

chlorine (0.5 ppm) administered at two-hour intervals once a week can reduce infection rates in 

Rainbow Trout by 63-73% without causing harm to the fish (Markiw 1992). Supply water 

treated with calcium cyanide (488 g/m2) mixed with chlorine gas (300 ppm) can be very 

effective at destroying M. cerebralis spores (Hoffman and Dunbar 1961). Water treated with 

chlorine (130-260 ppm) for 10 minutes may kill 100% of TAM spores present (Wagner 2002), 

and treating with chlorine (5,000 ppm) for 10 minutes is sufficient enough to destroy both 

triactinomyxon and myxospore (E. MacConnell, pers. comm. in Wagner 2002). Treating fry with 

chlorine (10 ppm) for 30 minutes may prevent whirling disease infection (Hoffman and 

O’Grodnick 1977). At the Roaring Judy Hatchery, a project is underway to install an ultraviolet 

system that kills M. cerebralis spores (CDW 2011). Treating water with 2537Å UV at doses of 

35mWs/cm2can be 86-100% effective at preventing whirling disease in Rainbow Trout fry 

(Hoffman 1974) and administering 1,300 mWs/cm2of UV under a static collimated beam, can 

inactivate 100% of the TAM spores present (Hedrick et al. 2000). Earthen pond substrate treated 

with quicklime (CaO) at concentrations greater than 380 g/m2for two weeks prior to introducing 

fish can prevent whirling disease infection by destroying M. cerebralis spores (Hoffman and 

Hoffman 1972).

Myxobolus cerebralis is frequently controlled by managing the intermediate host, Tubifex 

tubifex. Managers have observed that using concrete in aquaculture facilities can reduce the 

abundance of T. tubifex and thus limit the ability of M. cerebralis to reproduce (Mills et al. 1993, 

Ricciardi 2001). Maintaining water quality, reducing favorable habitat by preventing sediment 

accumulation in aquaculture (Crosier et al. 2012), and desiccating holding tanks, equipment, and 

intake pipes may help control T. tubifex (Kaster and Bushnell 1981). Lampricide TFM (3-

triflouromethyl-4-nitrophenol), administered at (4.2-14.0 mg/L) doses, is effective at destroying 

T. tubifex (Lieffers 1990). T. tubifex can also be treated in 30°C water for four days, causing 

triactinomyxon (TAM) spore production to stop, thus preventing the next stage of the parasites 

life cycle (El-Matbouli et al. 1999). T. tubifex ability to support M. cerebralis’ triactinomyxon 

(TAM) spore production may be due to genetic differences among T. tubifex populations. This 

variability may be an important factor in determining infection rates among fish (Baxa et al. 

2006) and therefore might support certain management practices (Stromberg 2006). There is 

evidence that electricity (1,000 s exposure to low-level DC voltage for 48 hrs) can destroy T. 

tubifex in aquaculture (R. Ingraham and T. Claxton, pers. comm. in Wagner 2002).
75

It has been proposed that selective processes are yielding a surviving population of fish that is 

more resistant to Myxobolus cerebralis infection on the Madison River, Montana (Vincent 2006). 

The implications of this for management are still unclear. However, research is continuing to 

evaluate the possibility of a developing resistance within salmonid populations (Stromberg 

2006).

It has been demonstrated that feeding Rainbow Trout with pellets containing (0.1%) Fumagillin 

is effective at reducing whirling disease infection. Two groups of Rainbow Trout were 

administered pellets from days 14-64 and 30-160 post infection. Approximately 10-20% of the 

medicated fish harbored spores, whereas 73-100% of non-medicated fish harbored spores (ElMatbouli and Hoffman 1991).

No effective treatments have been identified for Acineta nitocrae, Glugea hertwigi, 

Sphaeromyxa sevastopoli, or Trypanosoma acerinae.

4.9.4 Bacteria

Antimicrobial agents used to treat A. salmonicida infections include thiophenicol, furazolidone 

(Herman 1968), oxytetracycline (Herman 1968, Heo and Seo 1996, Wiklund and Dalsgaard 

1998), sulphamerazine, tetracycline, and a combination of trimethoprim (Heo and Seo 1996, 

Wiklund and Dalsgaard 1998) and sulphonamide (McCarthy and Roberts 1980). Others include 

flumequine (Michel et al. 1980), oxolinic acid (Hastings and McKay 1987, Heo and Seo 1996, 

Wiklund and Dalsgaard 1998), florfenicol (Inglis et al. 1991b), amoxycillin (Inglis et al. 1992), 

enrofloxacin (Stoffregen et al. 1993), chloramphenicol, neomycin, nitrofurantoin, and 

ciprofloxacin (Heo and Seo 1996, Wiklund and Dalsgaard 1998). Feed containing terramycin 

and romet are effective in treating A. salmonicida (MIDNR 2012). However, an increase in 

antimicrobial resistance was recognized in the United States beginning in 1967 (Wood 1967). 

Antimicrobial resistance by A. salmonicida has been discovered with the following agents: 

sulphonamides (Herman 1968), oxytetracycline (Tsoumas et al. 1989, Inglis et al. 1991a, Grant 

and Laidler 1993), a combination of sulphonamide and trimethoprim (Tsoumas et al. 1989, Grant 

and Laidler 1993), oxolinic acid (Hastings and McKay 1987, Tsoumas et al. 1989, Inglis et al. 

1991a, Höie 1992, Grant and Laidler 1993, Oppegaard and Sörum 1994), flumequin (Höie 

1992), and amoxycillin (Grant and Laidler 1993). The United States FDA approved broadspectrum, in-feed antibiotic AQUAFLOR® is now available to control mortality in finfish due to 

furunculosis (MAA 2012). Antimicrobial agents used to treat Renibacterium salmoninarum

include nitrofurans, bacitracin, chlortetracycline, oxytetracycline, novobiocin (Wolf and Dunbar 

1959), cephalosporins, gentamicin, clindamycin, lincomycin, oleandomycin, kitasamycin, 

spiramycin, penicillin (Austin 1985), cefazolin, tiamulin (Bandín et al. 1991), cephradine, 

rifampicin, (Brown et al. 1990), tetracycline (Austin 1985, Bandín et al. 1991), chloramphenicol 

(Wolf and Dunbar 1959, Austin 1985), and erythromycin (Wolf and Dunbar 1959, Austin 1985, 

Bandín et al. 1991, Lee and Evelyn 1994). All commonly used aquaculture disinfectants are 

considered effective against Piscirickettsia spp. (Fryer et al. 1990, 1992, Corbeil and Crane
76

2009). Antimicrobial agents used to treat P. salmonis include oxolinic acid and flumequin 

(Guardabassi and Courvalin 2006, Todar 2008, Corbeil and Crane 2009). 

Disinfecting eggs with 100 mg/L of iodine for 15 minutes may not eliminate vertical 

transmission of Bacterial Kidney Disease (BKD) but can reduce the severity of the disease 

(Bullock et al. 1978). Treating eggs with iodine at 250 or 500 mg/L for 15-120 minutes is 

effective at eliminating Renibacterium salmoninarum. However, after treatment, variable 

numbers of R. salmoninarum cells have survived. This is due to cells within the cell aggregates 

never coming in contact with the iodine (Evelyn et al. 1984, 1986). salmoninarum is inactivated 

by free chlorine (≤0.05 mg/L), which can be used as a disinfectant and to treat intake and 

effluent (Pascho et al. 1995).

The U.S. FDA approved vaccine Furogen® (Aqua Health, LTD) administered to brood stock 

prior to spawning has proven to be very effective at reducing A. salmonicida prevalence 

(GLFHC 2006). Very good to excellent results controlling and preventing out-breaks of 

furunculosis have also been obtained using an autogenous vaccine, produced by 

Microtechnologies; this treatment was accompanied by an immune-enhancing feed administered 

for three weeks prior and three weeks post vaccination, as well as stock thinning to reduce 

overall stress (GLFHC 2006). Vaccines derived from inactivated Piscirickettsia cf. salmonis

bacteria are considered ineffective against the bacterium. However, vaccines being developed 

using recombinant DNA have promise in combating P. salmonis infections (Corbeil and Crane 

2009). Further research is required to understand the practical application of this therapy.

Administration of bacteriophage to infected fish may help control outbreaks. In a study that 

administered the bacteriophage HER 110 to A. salmonicida HER 1107 infected Brook Trout 

(Salvelinus fontinalis), A. salmonicida populations declined by six log units (base 10) in 3 d. 

Further tests within fish populations are necessary to better understand the implementations of 

this therapy (Imbeault et al. 2006).

While supplements of lactic acid bacteria (Carnobacterium spp.) given with fish feed do not 

protect against Aeromonas salmonicida infection (Gildberg et al. 1995), such probiotic 

supplementation (also A. hydrophila and Vibrio fluvialis) can decrease mortality in Atlantic 

salmon (Salmo salar) and rainbow trout (Oncorhynchus mykiss) infected with A. salmonicida

(Irianto and Austin 2002). Carnobacterium strain K1 colonizes the intestinal tract of rainbow 

trout and inhibits A. salmonicida growth (Jöborn et al. 1997). Short-term bathing of presmolt 

Atlantic Salmon infected with furunculosis with siderophore-producing Pseudomonas 

fluorescens is another successful biological control (Gram et al. 1999, Smith and Davey 1993). 

Bath treatments with V. alginolyticus (Austin et al. 1995) can also lead to a reduction in mortality 

(Verschuere et al. 2000). Renogen® can reduce mortality rates in Piscirickettsia cf. salmonis

infected Pacific salmon (Torenzo et al. 2005).
77

Erythromycin-enhanced feed administered at 100 mg/kg/day for 21 days (Wolf & Dunbar 1959) 

or for 10 days (Austin 1985) is believed to be the most effective treatment for bacterial kidney 

disease (Hirvela-koski 2005). Erythromycin phosphate is an effective chemoprophylaxis in prespawning adult brood fish. Subcutaneous injections of 11 mg/kg (Hirvela-koski 2005) or 20 

mg/kg (Gudmundsdóttir et al. 2000, Pascho et al. 1991) administered to fish entering trapping 

facilities and at 21-30 day intervals thereafter (Hirvela-Koski 2005) has reduced mortality in respawning Pacific salmon by 10-50% (Groman and Klonz 1983). However, this treatment does 

not effectively destroy the pathogen from all eggs (Brown et al. 1990). A study by Lee and 

Evelyn (1994) showed female Coho Salmon treated with 20 mg/kg erythromycin prior to 

spawning yielded no vertical transmission of BKD. Treatment success relies on careful timing of 

the injections in adult salmonids before spawning (Elliott et al. 1989).

Culling is effective at preventing horizontal transmission of Piscirickettsia cf. salmonis (Torenzo 

et al. 2005). Culling infected stock has reduced the prevalence of Renibacterium salmoninarum

in aquaculture (Gudmundsdóttir et al. 2000, Maule et al. 1996, Pascho et al. 1991).

4.9.5 Viruses

Establishment of quarantines, culling, and stock density reduction during the winter and spring 

are beneficial management practices to prevent the spread of viruses (CFSPH 2007). VHS should 

be reported to Area Veterinarians in Charge (AVIC) or state veterinarians immediately upon 

diagnosis or recognition of the disease. Fish health surveillance programs and fallowing are also 

useful methods of control (CFSPH 2007). The U.S. Fish and Wildlife Service recommend 

implementation of the International Hazard Analysis and Critical Control Point (HACCP) 

planning standard to prevent the spread of VHS (Bakal 2012).

Rhabdovirus carpio is inactivated by UV irradiation (254 nm), gamma irradiation (103 krads), 

heating to 60°C (140°F) for 30 minutes, and exposure to pH 12 for 10 minutes, or pH 3 for 3 

hours (CFSPH 2007, World Organization for Animal Health 2009). Exposure to VHS can be 

prevented through use of spring water, specific pathogen free (SPF) stock, and separate 

cultivation of salmonids and flatfish (CFSPH 2007). Multiple means of VHS control are 

available to fish hatchery managers, including treatment of water with UV light subtype C (280-

200 nm wavelength) irradiation and heat (>15°C) (McAllister 1990), exposure to pH levels 

lower than 2.5 or higher than 12.2, desiccation of tanks and equipment (CFSPH 2007), 

minimization of stressors, cessation of water flow to adjacent waterways, and establishment of 

quarantines (Warren 1983, CFSPH 2007). 

Disinfection of live wells and other equipment potentially contaminated with Largemouth Bass 

Virus (LMBV) or VHS can be accomplished with a 10% household bleach/water solution (i.e., 

100 ml of household bleach to 900 mL of water). Waste water should be discarded away from 

any water body. The VHS virus is sensitive to ether, chloroform, glycerol, formalin, iodophor, 

sodium hydroxide, and sodium hypochlorite, which can be used as disinfectants (McAllister 

1990, CFSPH 2007). Rhabdovirus carpio is susceptible to oxidizing agents like sodium dodecyl 

sulphate, non-ionic detergents, and lipid solvents. The virus is inactivated by formalin, chlorine, 

iodine, NaOH, banzalkonium chloride, alkyltoluene, chlorhexidine gluconate, and cresol (Ahne 
78

and Held 1980, Ahne 1982, Fijan 1999, CFSPH 2007, Kiryu et al. 2007). Methisoprinol may be 

useful by inhibiting replication of Spring Viremia of Carp Virus (SVCV) in vitro; but further 

testing under culture conditions is necessary (Siwicki et al. 2003).

Single-stranded and double-stranded RNA injections can provide Rhabdovirus carpio protection 

for up to three weeks (Masycheva et al. 1995, Aliken et al. 1996). No effective anti-viral agents 

or commercial vaccines exist (CFSPH 2007) for VHS. 

5. Discussion

Only a small fraction (28%) of the more than 180 nonindigenous species which have become 

established in the Great Lakes are regulated by name primarily as invasives under federal or state 

law; just 26 plants, 10 fishes, 5 fish pathogens, 6 mollusks, 3 waterfleas, and 1 mysid within this 

subset, laws are a patchwork with only four diseases plus Zebra Mussels regulated by all states. 

In addition to the federal regulations applicable to the noxious weed Sparganium glomeratum, 

the Great Lakes states regulate 26 species of nonindigenous aquatic plants that are established in 

the basin. Seventy-three percent of the listed plants have documented moderate to high 

environmental and or socioeconomic impacts. An overlapping 35% have documented benefits. 

No individual plant species is regulated in all eight Great Lakes states. Some species (e.g., Black 

Alder) restricted in one state are recommended in another.

Many nonindigenous fishes are managed primarily as beneficial species (commercial and sport 

fisheries or prey fishes supporting those fisheries). Just 10 species of nonindigenous fish are 

regulated primarily as invasives.

Five species of fish pathogens, including two bacteria (Bacterial Kidney Disease (BKD) and 

furunculosis), a myxosporean (whirling disease), and two viruses (VHS and LMBV) are 

explicitly targeted by regulation by the Great Lakes states. Four of these species (all but 

furunculosis) are considered to have high environmental impact to the Great Lakes. Furunculosis 

is targeted primarily because it is considered a threat to hatchery operations for rearing 

salmonids.

Six species of mollusks are state-regulated. Of these, only Zebra Mussels are regulated in all 

eight Great Lakes states, though Quagga Mussels are listed in federal regulation so are also 

effectively regulated in all eight states as well. This list includes a mix of species with high 

(Dreissenid mussels), moderate (Asian Clam and New Zealand Mudsnail) and unknown 

(mystery snails) impacts. 

Three species of established nonindigenous waterfleas as well as one mysid are regulated by 

name by select Great Lakes states. These include two high impact raptorial waterfleas 

(Bythotrephes and Cercopagis) and two relatively recent invaders whose impact we are as yet 

unable to assess (Daphni lumholtzi and Hemimysis anomala).
79

Unregulated species are not necessarily innocuous. Our previous assessments identified only 35 

of the unregulated species (30%) as being low impact for all three assessments (environmental, 

socioeconomic and beneficial impacts) and an additional 10 species (9%) as being beneficial 

(Ulva flexuosa, Acentria epemerella, Lupinus polyphyllus, Mentha gracilis, Mentha spicata, 

Polygonum persicaria, Puccinellia distans, Salix alba, Salix fragilis, and Salix purpurea)

(Sturtevant et al. 2014). Most of the unregulated species (57%) are ‘understudied’ with little to 

no information on their impacts available in the literature. However, this list includes five species 

(4%) with known high impact: (Nitellopsis obtusa, Bithynia tentaculata, Ichthyocotylurus 

pileatus, Polygonum persicaria and Heterosporis) and 12 species (10%) with moderate impacts 

(Actinocyclus normanii fl. Subsalsa, Cylindrospermopsis raciborskii, Stephanodiscus 

binderanus, Ulva flexuosa, Ulva intestinalis, Carex acutiformix, Juncus compressus, Juncus 

gerardii, Juncus inflexus, Salix fragilis, Echinogammarus ischnus, and Piscirickettsia cf. 

salmonis). 

A growing body of literature provides a toolbox for control of nonindigenous species. However, 

most control efforts are successful at eradication only when the population is confined (e.g.,

aquaculture or other facilities, small ponds, limited geographic areas, etc). Control options 

suitable for established populations in open waters are extremely limited. Most control efforts 

focus on (1) controlling populations in locations where they directly impact human activities 

(e.g., aquaculture facilities and water intakes), (2) reducing populations to a level that minimizes 

impact to human socioeconomic endeavors (e.g., limiting impact to fisheries, agriculture, beach 

use, etc), (3) eradicating small, new infestations in confined geographic areas (e.g., a new 

infestation of Iris pseudacorus at a small pond), or (4) controlling spread along a specific 

pathway (e.g., ballast water treatment; using rotenone to poison fish in a canal).

Control options for many species remain extremely limited, but new technologies are emerging,

and the toolbox is growing. Options for control of established nonindigenous algae in open 

waters are limited. Only the largest two species (Ulva prolifera and Nitellopsis obtusa) can be 

effectively harvested mechanically, even then, they are often the first to recolonize (Pullman and 

Crawford 2010). Some species, particularly Cylindrospermopsis raciborskii, are associated with 

stratified water columns and mechanical destratification (mixing) may help prevent bloom 

development (Antenucci et al. 2005). Although in some cases, control options are extremely 

limited (e.g., to hand pulling) and large, established infestations may be difficult to control. 

Control options are at least available for all the established nonindigenous plants – with a broad 

array of herbicides and physical controls, as well as biological controls for some species, readily 

available. An array of options is also available for control of fishes – from harvesting and 

trapping to physical barriers, chemical piscicides, food web manipulations, and emerging genetic 

control technologies. Nonindigenous aquatic insects are currently controlled primarily via 

control of their host plants – a strategy also employed for a number of smaller invertebrates. 

Biocides exist for many additional species, though application in open systems remains 

unfeasible for most.
80

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7. Acknowledgements

Many thanks are given to the GLANSIS expert review panel for valuable input and feedback, 

including Anthony Ricciardi, Patricia Chow-Fraser, Hugh MacIsaac, Eugene Stoermer, Sarah 

Bailey, Hunter Carrick, Susan Galatowitsch, Jeff Gunderson, Rex Lowe, Nicholas Mandrak, and 

Robin Scribailo. We also appreciate the feedback received on individual species assessments 

from several external reviewers, including Steve Hensler, Tim Campbell, Titus Selheimer, Kevin 

Irons, Blake Ruebush, and Lisa Huberty. We thank the staff of the Cooperative Institute for 
130

Limnology and Ecosystem Research (CILER) and the Great Lakes Environmental Research 

Laboratory (GLERL) for providing many forms of support throughout this project. We also 

recognize past GLANSIS team members, including Gabriela Núñez, Kyle Dettloff, Emily Baker, 

Julie Larson, Mary McCarthy, Alex Bogdanoff and Katie Thompson, for their contributions and 

dedication to the improvement of GLANSIS. This research was funded through Great Lakes 

Restoration Initiative interagency agreement #DW-13-92312201-0. This is GLERL Contribution 

No. 1795.
131

Appendix A. Species Management Profiles

Notes: 

 Check federal, state/provincial, and local regulations for the most up-to-date information. 

 Check state and local regulations for the most up-to-date information regarding permits for 

control methods.

 Follow all label instructions.

A.1 Algae

Actinocyclus normanii f. subsalsa (Juhlin-Dannfelt) Hustedt, 1957

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Actinocyclus normanii f. subsalsa is one of the most pollution tolerant algal species and thrives 

in warm, shallow, and eutrophic waters (Edlund et al. 2000). The reduction of pollution and 

nutrient run-off would decrease the viable habitat for A. normanii f. subsalsa.

Bangia atropurpurea (Roth) Agardh, 1824

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The distribution of B. atropurpurea in the Great Lakes is associated with elevated salinity and 

eutrophic conditions (Lin and Blum 1976, Sheath and Cole 1984, Graham and Graham 1987, 

Jackson 1988, Stewart 2008). The reduction of pollution and nutrient run-off could decrease the 

viable habitat for B. atropurpurea.
132

Chaetoceros muelleri subsalsum J. R. Johansen and Rushforth, 1985

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The reduction of nutrient pollution, specifically of NaCl, would eliminate viable water conditions 

for C. muelleri var. subsalsum in the Great Lakes.

Chroodactylon ornatum (C. Agardh) Basson, 1979

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Contricribra guillardii (Hasle) K. Stachura-Suchoples & D.M. Williams

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species

Physical

There are no known physical control methods for this species

Chemical

Contricribra guillardii is a brackish water species. Therefore, the reduction of pollution and 

nutrient run-off could decrease the viable habitat for C. guillardii.

Cyclotella atomus Hustedt, 1937

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
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Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The reduction of pollution and nutrient run-off would decrease the viable habitat for C. atomus.

Cyclotella cryptica Reimann, Lewin, and Guillard, 1963

Regulations (pertaining to the Great Lakes region)Regulations (pertaining to the Great Lakes 

region)

There are no known regulations for this species

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Cyclotella cryptica is a euryhaline species (Liu and Hellebust 1976). As a result, reduction in 

run-off from winter road salt could decrease the chloride levels in the rivermouth areas and 

reduce the viable habitat for C. cryptica.

Cylindrospermopsis raciborskii (Wolosz.) Seenayya and Subbaraju, 1972

Regulations

There are no known regulations for this species.

Control

Biological

There are no known organisms that can degrade cylindrospermopsin. However, several studies 

have found that a variety of unidentified bacteria degraded 100% of saxitoxin, another toxin 

produced by C. raciborskii (Donovan et al. 2008, Ho et al. 2012).

Physical

Cylindrospermopsis raciborskii blooms are often associated with a stratified water column, 

which is one of the reasons they are so prevalent in shallow water bodies with a long turnover 

period. A mechanical system can be used to create artificial destratification to increase vertical 

mixing, introduce oxygen, and reduce internal nutrient loading. The installation of a 

destratification mechanism was attempted in a reservoir in Australia. While there was a reduction 
134

in internal nutrient loading, the increased turbidity of the water yielded a competitive advantage 

for C. raciborskii, which can move throughout the water column to compete for light (Atenucci 

et al. 2005).

Cylindrospermopsin can be absorbed by activated carbon with high mesopor capacity, and 

nanofiltration may be another viable option, but not enough research has been done to be 

conclusive about the efficacy of either technique. Saxitoxins can be absorbed by activated 

carbons that have a large faction of the pores being smaller than 1 nm (Westrick 2010).

Chemical

Cylindrospermopsin can be inactivated by chlorine, ozone, and hydroxyl radical treatments. 

Saxitoxin can be inactivated by chlorine (Westrick 2010). However, the use of copper-based 

algicides may inhibit the degradation of cylindrospermopsin (Smith et al. 2008).

Diatoma ehrenbergii Kützing, 1844

Regulations (pertaining to the Great Lakes region)

There are no known regulations of this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Diatoma ehrenbergii was recorded in eutrophic waters in the Great Lakes (Stoermer and Yang 

1969). The reduction of pollution and nutrient run-off could decrease the viable habitat for D. 

ehrenbergii.

Discostella pseudostelligera (Hustedt) Houk and Klee, 1939

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species

Control

Biological

There are no known biological control methods for this species

Physical

There are no known physical control methods for this species.

Chemical

Reduction of pollution and nutrient run-off would decrease the viable habitat for D. 

pseudostelligera.
135

Discostella woltereckii Hustedt, 1942

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Hymenomonas roseola Stein, 1878

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Nitellopsis obtusa (Desvaux in Loiseleur) J. Groves, (1919)

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species

Physical

It is difficult to mechanically remove all of a N. obtusa population from an inland lake because 

of the large amounts of biomass. Additionally, even when entire plants are removed, N. obtusa is 

typically the first macrophyte to reestablish the disturbed area because it is such an aggressive 

and efficient recolonizer (Pullman and Crawford 2010).

Chemical

Nitellopsis obtusa is very sensitive to common algaecides containing copper and endothall based 

compounds. When N. obtusa is still low growing, algaecide treatment can treat the entire 

organism. However, in taller individuals, the algaecide is absorbed in the top of the plant, killing 

that portion but leaving the bottom of the plant alive. This type of treatment has been found to be 

somewhat successful and is called a “hair cut treatment” by managers. The timing of the 

algaecide treatment is also important. Treatment early in the spring could help open up spawning 

habitat for native fish species, but N. obtusa or other nonnative aquatic plants are likely to 

recolonize these areas in the early summer (Pullman and Crawford 2010). Some applicators 

report control of Nitellopsis obtusa with 150ppb flumioxazin. 

Pleurosira laevis (Ehrenberg) Compère, (1843) 1982

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
136

Control

There are no known biological, physical or chemical methods of control for this species.

Skeletonema potamos (Weber) Hasle in Hasle & Evensen, (1970)

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Skeletonema subsalsum (Cleve-Euler) Bethge, (1912) 1928

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Sphacelaria fluviatilis Jao, 1943

Regulations (pertaining to the Great Lakes region)

There are no known methods of regulation for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Sphacelaria lacustris Schloesser and Blum, 1980

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Stephanodiscus binderanus Krieger, 1927

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.
137

Chemical

Stephanodiscus binderanus thrives in eutrophic waters. The reduction of pollution and nutrient 

run-off could decrease the viable habitat for S. binderanus.

Stephanodiscus subtilis (Van Goor) A. Cleve, 1951

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Stephanodiscus subtilis occurs primarily in eutrophic waters with elevated chloride levels (Millie 

and Lowe 1983). The reduction of pollution and nutrient run-off could decrease the viable 

habitat for S. subtilis.

Thalassiosira baltica Ostenfeld, 1901

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Thalassiosira lacustris (Grunow) Hasle in Hasle and Fryxell, 1977

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The ability of T. lacustris to reproduce and grow is highly dependent upon salinity. The 

reduction of pollution and run-off could decrease the viable habitat for this nonindigenous 

species.
138

Thalassiosira pseudonana (Hustedt) Hasle and Heimdal, (1957) 1970

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical methods of control for this species.

Thalassiosira weissflogii (Grunow) G. Fryxell & Hasle, (1896) 1977

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Thalassiosira weissflogii struggles to grow or reproduce at salinities below 5 ‰ (Stoermer 

1978). The reduction of pollution from road salt run-off could decrease the viable habitat for T. 

weissflogii.

Ulva (Enteromorpha) flexuosa subsp. flexuosa and flexuosa subsp. paradoxa (Wolfen 

ex Roth) J. Agardh, 1883

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The presence of U. flexuosa is associated with high nutrient availability and high levels of 

salinity (Shories et al. 1997, Lougheed and Stevenson 2004). The reduction of pollution and 

nutrient run-off could decrease the viable habitat for U. flexuosa.
139

Ulva (Enteromorpha) intestinalis Linnaeus, 1753

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

In locations outside of the Great Lakes, the distribution and abundance of U. intestinalis is 

dependent on salinity and nutrient levels (Kamer and Fong 2000, 2001, Messyasz and Rybak 

2011). The reduction of pollution and nutrient run-off could decrease the viable habitat for U. 

intestinalis.

Ulva (Enteromorpha) prolifera O.F. Müller, 1778

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

Ulva prolifera can be physically harvested from the water and beaches. However, depending on 

the size of the bloom this may not be an economically viable option (Ye et al. 2011).

Chemical

Ulva prolifera blooms occur in eutrophic marine waters. As a result, the reduction of pollution 

and nutrient run-off could decrease the viable habitat for U. prolifera.

A.2 Annelids

Branchiura sowerbyi

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Branchiura sowerbyi has not received much attention regarding control studies. The effects of 

industrial toxicants on Tubificidae species and explorations of their value as an indicator of 

environmental quality have been explored, but chemicals and heavy metals are not viable 
140

methods of control because of unknown and adverse effects on the surrounding environment 

(Das and Das 2005, Saha et al. 2006). However, there has been investigation into the control of 

Branchiura sowerbyi as a host of haemorrhagic thelohanellosis, which negatively impacts fishes

in aquaculture (Liyanage et al. 2003).

Biological

Brown Trout, Salmo trutta L., has been shown to prey on oligochaetes; its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, Brown Trout is itself an invasive species in the Great Lakes region and across nearly 

all of the United States.

Physical

Researchers found that Branchiura sowerbyi, which is a vector in transmission of Thelohanellus 

hovorkai (myxozoa) to fish, prefers muddy substrate, while other benthic oligochaetes that are 

not susceptible to myxozoa prefer sandy substrate, and suggested that replacing bottom substrate 

from mud to sand would lead to a shift in oligochaete communities from Branchiura sowerbyi to 

non-susceptible oligochaetes such as Limnodrilus socialus, therefore reducing disease in 

aquaculture fauna (Liyanage et al. 2003). 

Chemical

While there are no known chemical controls specifically for Branchiura sowerbyi, declines in 

Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in correlation 

with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of excess 

nutrients would help to reduce oligochaete populations.

Gianius aquaedulcis

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Little research has been pursued concerning control of Gianius aquaedulcis, likely because of its 

limited known range to the Niagara River in North America (Spencer and Hudson 2003).

Biological

While no there are no known biological controls specifically for Gianius aquaedulcis, Brown 

Trout, Salmo trutta L., has been shown to prey on oligochaetes, and its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, Brown Trout is itself nonindigenous to the Great Lakes region and across nearly all of 

the United States.

Research on benthic macroinvertebrate communities in southwestern Lake Ontario before and 

after the invasion of Dreissena polymorpha (Zebra Mussels) and Dreissena bugensis (Quagga 

Mussels) suggests that the presence of Dreissena helps to improve benthic habitat, facilitating 

increases in macroinvertebrates, including the tubificids Potamothrix vejdovskyi and 

Spirosperma ferox (Stewart and Haynes 1994). This indicates that control of invasive quagga and 
141

Zebra Mussels could facilitate improved control of benthic macroinvertebrates such as the 

tubificids.

Physical

There are no known physical control methods for this species.

Chemical

While there are no known chemical controls specifically for Gianius aquaedulcis, declines in 

Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in correlation 

with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of excess 

nutrients would help to reduce oligochaete populations.

Potamothrix bedoti (Piquet, 1913)

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

While no there are no known biological controls specifically for Potamothrix bedoti, Brown 

Trout, Salmo trutta L., has been shown to prey on oligochaetes, and its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, Brown Trout is itself nonindigenous to the Great Lakes region and across nearly all of 

the United States.

Research on benthic macroinvertebrate communities in southwestern Lake Ontario before and 

after the invasion of Dreissena polymorpha (Zebra Mussels) and Dreissena bugensis (Quagga 

Mussels) suggests that the presence of Dreissena helps to improve benthic habitat, facilitating 

increases in macroinvertebrates, including the tubificids Potamothrix vejdovskyi and 

Spirosperma ferox (Stewart and Haynes 1994). This indicates that control of invasive quagga and 

Zebra Mussels could facilitate improved control of benthic macroinvertebrates such as the 

tubificids.

Physical

Potamothrix bedoti has been shown to be more likely to occur in substrate with a high clay and 

silt content, but information on its ability to survive in other substrates is not available (Sauter 

and Gude 1996). However, this does indicate that substrate type is a possible physical control 

method to be further explored.

Chemical

While there are no known chemical controls specifically for Potamothrix bedoti, declines in 

Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in correlation 

with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of excess 

nutrients would help to reduce oligochaete populations.

Potamothrix moldaviensis Vejdovsky and Mrazek, 1902
142

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

While no there are no known biological controls specifically for Potamothrix moldaviensis, 

Brown Trout, Salmo trutta L., has been shown to prey on oligochaetes, and its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, Brown Trout is itself nonindigenous to the Great Lakes region and across nearly all of 

the United States.

Research on benthic macroinvertebrate communities in southwestern Lake Ontario before and 

after the invasion of Dreissena polymorpha (Zebra Mussels) and Dreissena bugensis (Quagga 

Mussels) suggests that the presence of Dreissena helps to improve benthic habitat, facilitating 

increases in macroinvertebrates, including the tubificids Potamothrix vejdovskyi and 

Spirosperma ferox (Stewart and Haynes 1994). This indicates that control of invasive quagga and 

Zebra Mussels could facilitate improved control of benthic macroinvertebrates such as the 

tubificids.

Physical

P. moldaviensis has been shown to be more likely to occur in sandy substrate with a clay and silt 

content of less than 10%, but information on its ability to survive in other substrates is not 

available (Sauter and Gude 1996). However, this does indicate that substrate type is a possible 

physical control method to be further explored.

Chemical

While there are no known chemical controls specifically for P. moldaviensis, declines in 

Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in correlation 

with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of excess 

nutrients would help to reduce oligochaete populations.

Potamothrix vejdovskyi Hrabe, 1941

Regulations (pertaining to the Great Lakes region)

Regulations

There are no known regulations for this species.

Note: Check federal, state/provincial, and local regulations for the most up-to-date information.

Control

Biological

While no there are no known biological controls specifically for Potamothrix vejdovskyi, Brown 

trout, Salmo trutta L., has been shown to prey on oligochaetes, and its removal from an 

experimental environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). 

However, brown trout is itself nonindigenous to the Great Lakes region and across nearly all of 

the United States.
143

Research on benthic macroinvertebrate communities in southwestern Lake Ontario before and 

after the invasion of Dreissena polymorpha (zebra mussels) and Dreissena bugensis (quagga 

mussels) suggests that the presence of Dreissena helps to improve benthic habitat, facilitating 

increases in macroinvertebrates, including the tubificids Potamothrix vejdovskyi and 

Spirosperma ferox (Stewart and Haynes 1994). This indicates that control of invasive quagga and 

zebra mussels could facilitate improved control of benthic macroinvertebrates such as the 

tubificids.

Physical

There are no known physical control methods for this species.

Chemical

While there are no known chemical controls specifically for Potamothrix vejdovskyi, declines in 

Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in correlation 

with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of excess 

nutrients would help to reduce oligochaete populations.

Ripistes parasita

Regulations

There are no known regulations for this species.

Control

Biological

While no there are no known biological controls specifically for Ripistes parasita, Brown trout, 

Salmo trutta L., has been shown to prey on oligochaetes, and its removal from an experimental 

environment led to rapid multiplication of benthic fauna (Wahab et al. 1989). However, brown 

trout is itself an invasive species in the Great Lakes region and across nearly all of the United 

States.

Physical

There are no known physical control methods for this species.

Chemical

Ripistes parasite has been found to occur in greater numbers where water quality is impaired by 

industrial pollution, therefore greater measures to control pollutants such as heavy metals and 

particulate matter might help control this oligochaete (Simpson and Abele 1984). Furthermore,

declines in Oligochaeta in southern Lake Michigan were recorded between 1980 and 1993 in 

correlation with reductions in phosphorus loads (Nalepa et al. 1998), suggesting that reduction of 

excess nutrients would help to reduce oligochaete populations.

A.3 Bacteria

Aeromonas salmonicida Emmerich and Weible, 1890
144

Regulations (pertaining to the Great Lakes)

Ohio requires source facilities outside the Great Lakes basin to document annual health 

inspections showing no furunculosis occurrences for the previous five years prior to importing 

salmonids to the Lake Erie watershed (Baird 2005). Indiana requires source facilities outside the 

basin to document they are furunculosis free prior to importing salmonid stock. Salmonids found 

carrying the pathogen, but asymptomatic, can be sold in state if source facilities are within the 

Great Lakes basin (Baird 2005). Michigan, Illinois, Wisconsin, and Minnesota also require 

imported salmonid health inspections. However, A. salmonicida is not a targeted pathogen. 

Minnesota allows the importation of furunculosis infected eggs, if prior egg treatments are 

approved (Baird 2005).

New York, Pennsylvania, Michigan, Illinois, Wisconsin, and Minnesota have instated similar 

baitfish regulations to control the spread of furunculosis and other fish pathogens. Those of New 

York include that bait harvested from inland waters for personal use is only permitted to be used 

within the same body of water from which it was taken and cannot be transported overland (with 

the exception of smelt, suckers, alewives, and blueback herring). Once transported, baitfish 

cannot be replaced to its original body of water (NYSDEC 2012).

Live or frozen bait harvested from inland New York waters for commercial purposes is only 

permitted to be sold or possessed on the same body of water from which it was taken and cannot 

be transported over land unless under a permit and or accompanied by a fish health certification 

report. Bait that is preserved and packaged by any method other than freezing, such as salting, 

can be sold and used wherever the use of bait fish is legal as long as the package is labeled with 

the name of the packager-processor, the name of the fish species, the quantity of fish packaged, 

and the means of preservation (NYSDEC 2012).

Certified bait may be sold for retail and transported overland as long as the consumer maintains a 

copy of a sales receipt that contains the name of the selling vendor, date sold, species of fish 

sold, and quantity of fish sold. Bait that has not been certified may still be sold but the consumer 

must maintain a sales receipt containing the body of water where the bait fish was collected and 

a warning that the bait cannot be transported by motor vehicle. Bait sold for resale require a fish 

health certification along with a receipt that contains the name of the selling vendor, date sold, 

species of fish sold, and quantity of fish sold, which must be kept for 30 days or until all bait is 

sold (NYSDEC 2012).

In addition to baitfish protections, prior to placing fish in New York waters, a fish health 

certification report must document that the fish are furunculosis free.

Control

Biological

While supplements of lactic acid bacteria (Carnobacterium spp.) given with fish feed do not 

protect against A. salmonicida infection (Gildberg et al. 1995), such probiotic supplementation 

(also A. hydrophila and Vibrio fluvialis) can decrease mortality in Atlantic Salmon (Salmo salar) 

and rainbow trout (Oncorhynchus mykiss) infected with A. salmonicida (Irianto and Austin 

2002). Carnobacterium strain K1 colonizes the intestinal tract of rainbow trout and inhibits A. 

salmonicida growth (Jöborn et al. 1997). Short-term bathing of presmolt Atlantic Salmon 
145

infected with furunculosis with siderophore-producing Pseudomonas fluorescens is another 

successful biological control (Gram et al. 1999, Smith and Davey 1993). Bath treatments with V. 

alginolyticus (Austin et al. 1995) can also lead to a reduction in mortality (Verschuere et al. 

2000).

Administration of bacteriophage to infected fish may also help control outbreaks. In a study that 

administered the bacteriophage HER 110 to A. salmonicida HER 1107 infected Brook Trout 

(Salvelinus fontinalis), A. salmonicida populations declined by six log units (base 10) in 3 d. 

Further tests within fish populations are necessary to better understand the implementations of 

this alternative therapy (Imbeault et al. 2006).

The US Food and Drug Administration approved vaccine Furogen® (Aqua Health, LTD) 

administered to brood stock prior to spawning has proven to be very effective at reducing A. 

salmonicida prevalence (GLFHC 2006). Very good to excellent results controlling and 

preventing out-breaks of furunculosis have also been obtained using an autogenous vaccine, 

produced by Microtechnologies; this treatment was accompanied by an immune-enhancing feed 

administered for three weeks prior and three weeks post vaccination, as well as stock thinning to 

reduce overall stress (GLFHC 2006).

Physical

There are no known physical control methods for this species.

Chemical

Antimicrobial agents used to treat A. salmonicida infections include thiophenicol, furazolidone 

(Herman 1968), oxytetracycline (Herman 1968, Heo and Seo 1996, Wiklund and Dalsgaard 

1998), sulphamerazine, tetracycline and a combination of trimethoprim (Heo and Seo 1996, 

Wiklund and Dalsgaard 1998) and sulphonamide (McCarthy and Roberts 1980). Others include 

flumequine (Michel et al. 1980), oxolinic acid (Hastings and McKay 1987, Heo and Seo 1996, 

Wiklund and Dalsgaard 1998), florfenicol (Inglis et al. 1991b), amoxycillin (Inglis et al. 1992), 

enrofloxacin (Stoffregen et al. 1993), chloramphenicol, neomycin, nitrofurantoin and 

ciprofloxacin (Heo and Seo 1996, Wiklund and Dalsgaard 1998). Feed containing terramycin 

and romet are effective in treating A. salmonicida (MIDNR 2012).

However, an increase in antimicrobial resistance was recognized in the United States beginning 

in 1967 (Wood 1967). Antimicrobial resistance by A. salmonicida has been discovered with the 

following agents: sulphonamides (Herman 1968), oxytetracycline (Tsoumas et al. 1989, Inglis et 

al. 1991a, Grant and Laidler 1993), a combination of sulphonamide and trimethoprim (Tsoumas 

et al. 1989, Grant and Laidler 1993), oxolinic acid (Hastings and McKay 1987, Tsoumas et al. 

1989, Inglis et al. 1991a, Höie 1992, Grant and Laidler 1993, Oppegaard and Sörum 1994), 

flumequin (Höie 1992), and amoxycillin (Grant and Laidler 1993).The United States Food and 

Drug Administration approved broad-spectrum, in-feed antibiotic AQUAFLOR® is now 

available to control mortality in finfish due to furunculosis (MAA 2012).

Other

Minimizing fish stress can reduce the risk of disease outbreak (FTS 2012).
146

Piscirickettsia cf. salmonis

Regulations (pertaining to the Great Lakes)

There are no known regulations for this species.

Control

Biological

Vaccines derived from inactivated P. cf. salmonis bacterins are considered ineffective against the 

bacterium. However, vaccines being developed using recombinant DNA have promise in 

combating P. salmonis infections (Corbeil and Crane 2009). Further research is required to 

understand the practical application of this therapy.

Physical

Culling is effective at preventing horizontal transmission of P. cf. salmonis (Torenzo et al. 2005).

Chemical

Renogen® can reduce mortality rates in P. cf. salmonis infected Pacific salmon (Torenzo et al. 

2005). All commonly used aquaculture disinfectants are considered effective against

Piscirickettsia spp. (Fryer et al. 1990 and 1992, Corbeil and Crane 2009,). Antimicrobial agents 

used to treat P. salmonis include oxolinic acid and flumequin (Guardabassi and Courvalin 2006, 

Todar 2008, Corbeil and Crane 2009).

Other

Establishment of muskellunge fingerling index surveys may help monitor trends in spawning 

success and or fingerling survival (Thomas and Faisal 2009).

Renibacterium (Corynebacterium) salmoninarum Sanders and Fryer, 1980

Regulations (pertaining to the Great Lakes)

Ohio requires source facilities outside the Great Lakes basin to document annual health 

inspections showing no bacterial kidnsey disease (BKD) occurrences for the previous five years 

prior to importing salmonids to the Lake Erie watershed (Baird 2005). Indiana requires source 

facilities outside the basin to document they are BKD free prior to importing salmonid stock. 

Asymptomatic salmonids found carrying the pathogen can be sold in-state if source facilities are 

within the Great Lakes basin (Baird 2005). Michigan, Illinois, Wisconsin, and Minnesota also 

require imported salmonid health inspections (NCRAC 2010ab). Minnesota allows the 

importation of infected eggs, if prior egg treatments have been approved (Baird 2005).

New York, Pennsylvania, Ohio, Michigan, Indiana, Illinois, Wisconsin, and Minnesota have 

instated similar baitfish regulations to control the spread of BKD and other fish pathogens. Those 

of New York include that bait harvested from inland waters for personal use is only permitted to 

be used within the same body of water from which it was taken and cannot be transported 

overland (with the exception of smelt, suckers, Alewives, and Blueback Herring). Once 

transported, baitfish cannot be replaced to its original body of water (NYSDEC 2012a).

Live or frozen bait harvested from inland New York waters for commercial purposes is only 

permitted to be sold or possessed on the same body of water from which it was taken and cannot 
147

be transported over land unless under a permit and or accompanied by a fish health certification 

report. Bait that is preserved and packaged by any method other than freezing, such as salting, 

can be sold and used wherever the use of baitfish is legal as long as the package is labeled with 

the name of the packager-processor, the name of the fish species, the quantity of fish packaged, 

and the means of preservation (NYSDEC 2012a).

Certified bait may be sold for retail and transported overland as long as the consumer maintains a 

copy of a sales receipt that contains the name of the selling vendor, date sold, species of fish 

sold, and quantity of fish sold. Bait that has not been certified may still be sold but the consumer 

must maintain a sales receipt containing the body of water where the baitfish was collected and a 

warning that the bait cannot be transported by motor vehicle. Bait sold for resale require a fish 

health certification along with a receipt that contains the name of the selling vendor, date sold, 

species of fish sold, and quantity of fish sold, which must be kept for 30 days or until all bait is 

sold (NYSDEC 2012).

In addition to baitfish protections, prior to placing fish in New York waters, a fish health 

certification report must document that the fish are BKD free.

Control

Biological

There are no known biological control methods for this species.

Physical

Culling infected stock has reduced the prevalence of R. salmoninarum in aquaculture 

(Gudmundsdóttir et al. 2000, Maule et al. 1996, Pascho et al. 1991).

Chemical

Antimicrobial agents used to treat R. salmoninarum include nitrofurans, bacitracin, 

chlortetracycline, oxytetracycline, novobiocin (Wolf and Dunbar 1959), cephalosporins, 

gentamicin, clindamycin, lincomycin, oleandomycin, kitasamycin, spiramycin, penicillin (Austin 

1985), cefazolin, tiamulin (Bandín et al. 1991), cephradine, rifampicin, (Brown et al. 1990), 

tetracycline (Austin 1985, Bandín et al. 1991), chloramphenicol (Austin 1985, Wolf and Dunbar 

1959), and erythromycin (Austin 1985, Bandín et al. 1991, Lee and Evelyn 1994, Wolf and 

Dunbar 1959).

Erythromycin-enhanced feed administered at 100 mg/kg/day for 21 days (Wolf & Dunbar 1959) 

or for 10 days (Austin 1985) is believed to be the most effective treatment for bacterial kidney 

disease (Hirvela-koski 2005). Erythromycin phosphate is an effective chemoprophylaxis in prespawning adult brood fish. Subcutaneous injections of 11 mg/kg (Hirvela-koski 2005) or 20 

mg/kg (Gudmundsdóttir et al. 2000, Pascho et al. 1991) administered to fish entering trapping 

facilities and at 21-30 day intervals thereafter (Hirvela-Koski 2005) has reduced mortality in respawning Pacific salmon by 10-50% (Groman and Klonz 1983). However, this treatment does 

not effectively destroy the pathogen from all eggs (Brown et al. 1990). A study by Lee and 

Evelyn (1994) showed female Coho Salmon treated with 20 mg/kg erythromycin prior to 

spawning yielded no vertical transmission of BKD. Treatment success relies on careful timing of 

the injections in adult salmonids before spawning (Elliott et al. 1989).
148

Disinfecting eggs with 100 mg/L of iodine for 15 minutes may not eliminate vertical 

transmission, but can reduce the severity of the disease (Bullock et al. 1978). Treating eggs with 

iodine at 250 or 500 mg/L for 15-120 minutes is effective at eliminating R. salmoninarum. 

However, after treatment, variable numbers of R. salmoninarum cells have survived. This is due 

to cells within the cell aggregates never coming in contact with the iodine (Evelyn et al. 1984, 

1986). Renibacterium salmoninarum is inactivated by free chlorine (≤0.05 mg/L), which can be 

used as a disinfectant and to treat intake and effluent (Pascho et al. 1995).

Other

Prevention is the preferred control method of BKD in cultured stocks (World Organization for 

Animal Health 2003).

A.4 Bryozoa

Lophopodella carteri (Hyatt, 1865)

Regulations

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

Lophopodella carteri colonies grow on solid substrata (Lauer et al. 1999), therefore, physical 

removal methods such as scraping may be viable for localized areas. 

Chemical

Chemical biocides have been used as anti-fouling agents to remove sessile macroinvertebrates 

from shipping equipment and industrial intakes, but none have been approved for use on 

bryozoans as of yet (United States 2011). The Great Lakes and Mississippi River Interbasin 

Study (GLMRIS 2012) suggests that alteration of water quality using carbon dioxide, ozone, 

nitrogen, and/or sodium thiosulfate could be effective in preventing upstream and downstream 

movement of bryozoans. 

Pardue and Wood (1980) determined baseline toxicity of four heavy metals to three species of 

phylactolaemate bryozoa (L. carteri, Pectinatella magnifica Leidy, and Plumatella emarginata 

Oka). They recorded 96-hr LC50 values (lethal concentration for 50 percent of organisms tested) 

of L. carteri, observing greatest sensitivity to cadmium (LC50 0.15 mg/L), followed by copper 

(LC50 0.51 mg/L), chromium (LC50 1.56 mg/L), and zinc (LC50 5.63 mg/L). It should be noted 

that the toxicity of these metals were not tested as control measures, but as a demonstration of 

the usefulness of some bryozoans as biomonitors of water quality. However, comparison of the 

96-hr LC50 data to toxicity data from other studies indicates that the bryozoa are more sensitive 

to the tested metals than many other invertebrates and fish, indicating potential as chemical 

controls with further research (Pardue and Wood 1980). 
149

A.5 Coelenterates

Cordylophora caspia (Pallas, 1771)

Regulations

There are no known regulations for this species.

Control

Control of Cordylophora caspia will likely focus on its potential role as a biofouling agent. 

Cordylophora spp. has been documented colonizing the inner walls of power plants in Europe 

and the United States (Folino 2000), primarily causing blockages in nozzles and tailpipes of 

rapid gravity filter beds (RGFs) (Mant et al. 2011). The menont life-stage of Cordylophora 

caspia often found in hydroelectric intakes, is both drought and temperature resistant which may 

prove an obstacle to control (Gutierre 2012).

Biological

In estuarine, brackish habitats such as San Francisco Bay on the Pacific and the North American 

Atlantic coast, nudibranchs such as Tenellia adspersa feed on Cordylophora caspia and other 

hydroids (Mills and Sommer 1995). However, unlike hydroids, nudibranchs are exclusively seadwelling invertebrates (Anderson 1995), and thus are only a source of biological control in 

brackish areas where the macroinvertebrates’ habitats intersect. 

Physical

Thermal treatments of >37°C have been proven effective in eradicating colonies of

Cordylophora spp. sampled from the walls of power plant intakes (Folino 2000), but are not 

efficient in water treatment facilities where there is no residual heat energy available (Mant et al. 

2011). 

Chemical

Gutierre (2012) found that Cordylophora caspia is completely eradicated at pH levels of 4.0 and 

10.0, with increasing survival rates in between, and suggested maintaining pH levels at 10.0 for 6 

hours or more by injection of NaOH to reduce and eliminate colonies. Chlorine treatments 

negatively affect Cordylophora growth, but treatments as high as 5 mg/L for periods of 105 

minutes have been unsuccessful in completely eradicating colonies (Mant et al. 2011). 

Furthermore, chlorine use is highly regulated at water treatment facilities where Cordylophora 

most frequently cause problems. Hydroids are sensitive to vanadium leeching from slag stones 

used in riverbank reinforcement, and are sensitive to heavy metals in general, especially 

mercury, copper, cadmium, and arsenic, though it is unlikely that these will be useful in control 

(Ringelband and Karbe 1996).

Craspedacusta sowerbyi Lankester, 1880

Regulations

There are no known regulations for this species.

Control
150

Craspedacusta sowerbyi has spread across temperate climates for more than a century, but 

despite experimental observation of its possible contribution to trophic cascade effects 

(Jankowski et al. 2005), and studies on predation habits (Dodson and Cooper 1983, Spadinger 

and Maier 1999, Dendy 1978), little research on control is available. 

Biological

There are no known biological control methods for this species. Craspedacusta sowerbyi 

populations are not checked by predation (Jankowski et al. 2005). 

Physical

There are no known physical control methods for this species. Hydrozoan hydromedusae blooms 

are known to be temperature dependent (Ma and Purcell 2005), but polyps and especially the 

overwintering podocysts are more resistant to varying physical conditions (Peard 2002).

Chemical

There are no known chemical control methods for this species. 

A.6 Amphipods

Echinogammarus ischnus (Stebbing, 1899)

Regulations

There are no known regulations for this species.

Control

Biological

Benthic invertebrates including Echinogammarus ischnus are a major part of the native yellow 

perch (Perca flavescens) diet (Gonzalez and Burkart 2004). Echinogammarus ischnus has also 

become prey of the invasive Round Goby, Neogobius melanostomus (Gonzalez and Burkart 

2004). The spread of dreissenid-covered substrate across the Great Lakes region has created an 

ideal habitat for Echinogammarus ischnus, where it is less susceptible to predation, indicating 

that efforts to control Dreissena spp. could also aid control of Echinogammarus ischnus

(Gonzalez and Burkart 2004). 

A parasitic water mold (oomycete) detected in the upper St. Lawrence river is likely responsible 

for reducing Echinogammarus ischnus abundance despite favorable physical and chemical 

conditions (Kestrup et al. 2011). The oomycete also infects the native amphipod Gammarus 

fasciatus, but its effects are significantly less severe than in Echinogammarus ischnus (Kestrup et 

al. 2011). 

Physical

Echinogammarus ischnus can tolerate a maximum water temperature between 31.0°C and 

32.2°C before irreversible physiological damage and mortality occur (Wijnhoven et al. 2003). 

Electron beam irradiation can be used to control microorganisms in aquatic pathways, including 

Echinogammarus ischnus (USACE 2012b). Electron beam irradiation is a non-selective control 
151

method which exposes water to low doses of radiation using gamma-sterilizers or electron 

accelerators, breaking down DNA in living organisms while leaving behind no by-products 

(USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms including E. 

ischnus in water treatment facilities and narrow channels, where UV filters can be used to emit 

UV light into passing water, penetrating cell walls and rearranging DNA of microorganisms 

(USACE 2012b).

Chemical

There are no known chemical control methods for this species

Gammarus tigrinus Sexton 1939

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Note: Check federal, state/provincial, and local regulations for the most up-to-date information.

Control

Biological

There are no known biological control methods for this species

Physical

Gammarus tigrinus can tolerate maximum water temperatures between 32.2°C and 34.2°C 

before irreversible physiological damage and mortality occur (Wijnhoven et al. 2003). In a study 

of the effects of tide gate operation on hypoxic conditions in the Back River and Savannah River 

estuaries in Savannah, Georgia, it was found that oxygen saturation levels below 30% are lethal 

to experimental populations of amphipods, and that Gammarus tigrinus is especially susceptible 

to low dissolved oxygen levels, exhibiting mortality within three hours of exposure to levels 

between 12 and 18% (Winn and Knott 1992).

Chemical

There are no known chemical control methods for this species.

A.7 Cladocerans

Bosmina coregoni Baird, 1857

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. 

Control

Biological

Young-of-year bloaters (Coregonus hoyi) have been shown to surface feed on Eubosmina 

coregoni in Lake Michigan, but their effectiveness as a control is unknown, especially because 

they have been documented moving to the benthos earlier to avoid competition from the invasive 

Alewife (Alosa pseudoharengus) (Crowder and Crawford 1984). Bythotrephes spp. are known to 

consume E. coregoni in Russia and the United States (Grigorovich et al. 1998), and E. coregoni
152

populations declined significantly immediately following the invasion of the waterflea 

Bythotrephes longimanus in the Laurentian Great Lakes Michigan, Huron, and Erie, with direct 

predation by B. longimanus the most likely explanation (Barbiero and Tuchman 2004). E. 

coregoni levels in B. longimanus-invaded areas have remained at low levels, indicating a 

significant population-wide impact, but the long-term effectiveness of B. longimanus as a 

biological control method is unknown (Barbiero and Tuchman 2004). 

Physical

E. coregoni may be transported over land by recreational boaters (Suchy and Hann 2007). 

Though it is not as likely to foul gear and attach to equipment as the more commonly known 

Spiny Waterflea (Bythotrephes longimanus) and Fishhook Waterflea (Cercopagis pengoi), the 

same responsible maintenance and cleaning methods are recommended to prevent spread 

between water bodies, including cleaning all aquatic equipment with high pressure water (>250 

psi) or hot water (>50°C) after each use (Ontario’s Invading Species Awareness Program). 

Electron beam irradiation and ultraviolet light treatments have been used to control Spiny and 

Fishhook Waterfleas in aquatic pathways, and are likely effective against E. coregoni (USACE 

2012b). Another possible non-selective pathway control is high water turbidity, which may 

decrease zooplankton (especially cladoceran) abundances due to the negative effects of 

suspended clay particles on filtering and assimilation rates (Suchy and Hann 2007). 

Chemical

E. coregoni is a freshwater cladoceran. Mortality has been documented quickly at salinities >3% 

(Nauwerck 1991). Gemza (1995) documented a shift from copepods to cladocerans as dominant 

zooplankton at increasingly eutrophic sites in Severn Sound, Lake Huron. Zooplankton biomass 

generally increases with increasing eutrophication, so reduction of excessive nutrient pollution 

causing abnormal eutrophication could help control E. coregoni (Gemza 1995). A study of the 

effects of cadmium and zinc on Lake Michigan zooplankton found that E. coregoni was 

significantly reduced by separate and combined treatments of 2 μg Cd/L and 100 μg Zn/L, with 

negative effects primarily due to zinc (Marshall et al. 1981). A more recent study on the effects 

of copper sulfate (used to control algal biomass in eutrophic water bodies) and Carbaryl (used to 

control aquatic pests) on zooplankton found that levels of 50 μg/L Cu and 20 μg/L Carbaryl 

individually reduced E. coregoni biomass by >50% (Havens 1994). 

Bythotrephes longimanus

Regulations (pertaining to the Great Lakes region)

In Wisconsin, the Spiny Waterflea is a prohibited invasive species (WI Administrative Code § 

NR 40.04), which indicates that it is likely to survive and spread if introduced into the state, 

potentially causing economic or environmental harm or harm to human health (WI

Administrative Code § NR 40.02). With certain exceptions, it is unlawful to transport, possess, 

transfer or introduce a prohibited invasive species in Wisconsin (WI Administrative Code § NR 

40.04). In Minnesota, the Spiny Waterflea is a regulated invasive species (MN Administrative

Rules § 6216.0260). It is legal to possess, sell, buy, and transport regulated invasive species, but 

no person may introduce a regulated invasive species without a permit (MN Administrative

Rules § 6216.0265 Subpart 1). 
153

Control

Like the confamilial Fishhook Waterflea (Cercopagis pengoi), the Spiny Waterflea is most likely 

to be spread on aquatic equipment, especially fishing lines. Consequently, public education is a 

significant method of control which can greatly reduce incidences of species transfer by unaware 

or incautious anglers (Jacobs and MacIsaac 2007, Lui et al. 2010). 

Biological

Bythotrephes longimanus is consumed by rainbow smelt, lake herring, lake hitefish, Yellow 

Perch, White Perch, White Bass, Walleye, Alewife, Bloater Chub, Emerald Shiner, Spottail 

Shiner, Deepwater Sculpin, and Chinook Salmon in the Great Lakes (Bur et al. 1986,

Makarewicz and Jones 1990, Branstrator and Lehman 1996). Bythotrephes longimanus’ 

defensive tailspine has been observed increasing in size throughout the summer in response to 

predation pressure (Straile and Halbich 2000). Consequently, larger fish are more likely to be 

successful predators (Branstrator and Lehman 1996). The opossum shrimp (Mysis relicta) has 

been observed eating B. longimanus in Ontario lakes, but the frequency of consumption appeared 

related to abundance of the invader and alternate prey (Nordin et al. 2008). 

Physical

Bythotrephes longimanus collects in gelatinous clumps on fishing lines, downrigger cables, and 

other aquatic equipment (Lui et al. 2010). Responsible maintenance and cleaning methods are 

recommended to prevent spread between water bodies, including cleaning all aquatic equipment 

with high pressure (>250 psi) or hot (>50°C) water after each use (Ontario’s Invading Species 

Awareness Program). The acute upper lethal temperature level for B. longimanus, at which death 

occurs rapidly, is 40°C (USACE 2012b), and a study found that B. longimanus specifically 

requires 10 minutes treatment with 43°C water to ensure 100% mortality (Beyer et al. 2011). 

Fishing lines designed specifically to prevent the spread of waterfleas, such as the Flea Flicker 

brand, have been proven effective in significantly reducing fouling on lines, indicating their 

importance as a management tool (Jacobs and MacIsaac 2007).

Electron beam irradiation has been used to control microorganisms in aquatic pathways, 

including Bythotrephes longimanus (USACE 2012b). Electron beam irradiation is a nonselective control method which exposes water to low doses of radiation using gamma-sterilizers 

or electron accelerators, breaking down DNA in living organisms while leaving behind no byproducts (USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms 

including B. longimanus in water treatment facilities and narrow channels, where UV filters can 

be used to emit UV light into passing water, penetrating cell walls and rearranging DNA of 

microorganisms (USACE 2012b).

Chemical

There are no known chemical control methods for this species

Cercopagis pengoi

Regulations (pertaining to the Great Lakes region)

In Wisconsin, the Fishhook Waterflea is a prohibited invasive species (WI Administrative Code 

§ NR 40.04), meaning that it is unlawful to transport, possess, transfer, or introduce the species 
154

within or into the state without a permit as defined under Wisconsin Administrative Code § NR 

40.06. 

Control

Cercopagis pengoi is most likely to be spread on aquatic equipment, especially fishing lines. 

Consequently, public education is a significant method of control which can greatly reduce 

incidences of species transfer by unaware or incautious anglers (Jacobs and MacIsaac 2007). 

Biological

Pothoven et al. (2007) found that adult large Alewives (Alosa pseudoharengus) (>100 mm) 

consume Cercopagis pengoi in Lake Michigan, but not significantly enough to control the 

species, concluding that the Alewife prefers Bythotrephes longimanus due to its larger size and 

conspicuousness. In contrast, a study of C. pengoi as a prey item in Lake Ontario found that at 

least 70% of Alewives larger than 70 mm contained C. pengoi spines (Bushnoe et al. 2003). The 

same study also found spines in Rainbow Smelt (Osmerus mordax) stomachs (Bushnoe et al. 

2003). Rainbow Smelt historically consume cladocerans in the Great Lakes, but prefer larger 

prey and may select Bythotrephes longimanus over C. pengoi where both occur (Pothoven et al. 

2009). Gorokhova et al. (2004) found that in the northern Baltic proper, Herring (Clupea 

harengus L.) and Sprat (Sprattus sprattus L.) are the dominant predators of C. pengoi, and a 

possible source of biological control through fisheries management, though it is possible that 

fully mature resting eggs may survive passage through fish digestive systems as has been 

observed with B. longimanus eggs in Yellow Perch (Perca flavescens). B. longimanus is also 

known to consume C. pengoi, but not as a main prey item (Cavaletto et al. 2010). 

Physical

Cleaning all aquatic/fishing equipment, including downrigger lines and monofilament on reels, is 

important in areas where this species is present. Responsible maintenance and cleaning methods 

are recommended to prevent spread between water bodies, including cleaning all aquatic 

equipment with high pressure (>250 psi) or hot (>50°C) water after each use (Ontario’s Invading 

Species Awareness Program). Bythotrephes longimanus has been documented spreading by 

transfer of diapausing eggs on fishing gear, which are more resilient than adult waterfleas 

(Jacobs and MacIsaac 2007). Because of its similar life history to B. longimanus, it is likely that 

Cercopagis pengoi can also be spread by introduction of diapausing eggs to previously 

uninvaded waters as well as by transfer of fully developed adult specimens (Jacobs and MacIsaac 

2007). Fishing lines designed specifically to prevent the spread of waterfleas, such as the Flea 

Flicker brand, have been proven effective in significantly reducing fouling on lines, indicating 

their importance as a management tool (Jacobs and MacIsaac 2007). 

Electron beam irradiation has been used to control microorganisms in aquatic pathways, 

including C. pengoi (USACE 2012b). Electron beam irradiation is a non-selective control 

method which exposes water to low doses of radiation using gamma-sterilizers or electron 

accelerators, breaking down DNA in living organisms while leaving behind no by-products 

(USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms, including 

C. pengoi, in water treatment facilities and narrow channels, where UV filters can be used to 

emit UV light into passing water, penetrating cell walls and rearranging DNA of microorganisms 

(USACE 2012b).
155

Chemical

There are no known chemical control methods for this species. 

Daphnia galeata galeata Sars, 1864

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There is little control information available on a species-specific level for Daphnia galeata 

galeata, especially as related to its presence in North America. However, many control methods 

used for more commonly known waterfleas may work for D. g. galeata as well. 

Biological

Various parasites have been shown to reduce host density and population survival in 

experimental Daphnia populations in Europe (Ebert 2005). However, it is unclear whether 

parasites regulate natural Daphnia populations, as all experiments have been completed under 

lab conditions. Research on D. g. galeata is lacking, but many invertebrates are likely predators 

of Daphnia spp. where they occur in Europe and North America, including Great Lakes species 

such as the predacious phantom midge Chaoborus flavicans, the waterfleas Leptodora spp. and 

Bythotrephes longimanus, (Lysebo 1995). Many small fish of species such as Yellow Perch

(Perca flavescens), Three-Spined Stickleback (Gasterosteus aculeatus), Alewife (Alosa 

pseudoharengus), Bluegill (Lepomis macrochirus), and ciscos (Coregonus spp.) have been 

documented consuming daphnids and other zooplankton in Canada and the United States (Mills 

and Forney 1983, Post and McQueen 1987, Hulsmann and Mehner 1997), but predation may be 

insufficient for control in most cases.

Physical

Electron beam irradiation has been used to control microorganisms in aquatic pathways, 

including D. g. galeata (USACE 2012b). Electron beam irradiation is a non-selective control 

method which exposes water to low doses of radiation using gamma-sterilizers or electron 

accelerators, breaking down DNA in living organisms while leaving behind no by-products 

(USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms including 

D. g. galeata in water treatment facilities and narrow channels, where UV filters can be used to 

emit UV light into passing water, penetrating cell walls and rearranging DNA of microorganisms 

(USACE 2012b). Hot water (e.g., boat washing) may be a viable method for killing adults, but 

resting stages are likely resistant.

Chemical

There are no known chemical control methods for this species.
156

Daphnia lumholtzi

Regulations (pertaining to the Great Lakes region)

In Wisconsin, Daphnia lumholtzi is a prohibited invasive species (WI Administrative Code § NR 

40.04), meaning that it is unlawful to transport, possess, transfer, or introduce the species within 

or into the state without a permit as defined under Wisconsin Administrative Code § NR 40.06. 

Control

Biological

Daphnia lumholtzi is preyed upon by a variety of zooplanktivorous fishes, including Inland 

Silversides (Menidia beryllina) in Lake Texoma (Oklahoma-Texas), Bluegill (Lepomis 

macrochirus), White Bass (Morone chrysops), White Crappie (Pomoxis annularis), and Black 

Crappie (Pomoxis nigromaculatus) in Lake Chautauqua, Illinois (Lienesch and Gophen 2001, 

Lemke et al. 2003). The degree to which these fishes may be able to control D. lumholtzi

populations is not certain. 

Physical

D. lumholtzi is likely transferred through anthropogenic vectors, including on recreational boats 

(Frisch et al. 2013). Its resting eggs (ephippia) feature a long point and hairs on the margin that 

serve as hooks, possibly aiding in attachment to boats or macrophytes caught on boats 

(Dzialowski et al. 2000). Therefore, the same precautions recommended to prevent the spread of 

more commonly known waterfleas should be taken to help prevent D. lumholtzi distribution, 

namely cleaning all aquatic/fishing/boating equipment, including downrigger lines and 

monofilament on reels, using high pressure (>250 psi) or hot (>50°C) water after each use 

(OFAH n.d.). 

While D. lumholtzi is not listed as a target species by the United States Army Corps of Engineers 

Great Lakes and Mississippi River Interbasin Study (USACE GLMRIS), the methods suggested 

to control Spiny and Fishhook Waterfleas in aquatic pathways would likely control D. lumholtzi 

as well. Electron beam irradiation has been used to control microorganisms in aquatic pathways 

by exposing water to low doses of radiation using gamma-sterilizers or electron accelerators, 

breaking down DNA in living organisms while leaving behind no by-products (USACE 2012b). 

Ultraviolet (UV) light can also effectively control microorganisms, in water treatment facilities 

and narrow channels, where UV filters can be used to emit UV light into passing water, 

penetrating cell walls and rearranging DNA of microorganisms (USACE 2012b).

Chemical

There are no known chemical control methods for this species.

Eubosmina maritima P.E. Müller, 1867

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Note: Check federal, state/provincial, and local regulations for the most up-to-date information.
157

Control

Biological

The mysid shrimps, Mysis mixta and Mysis relicta, consume Eubosmina maritima diapausing 

eggs (ephippia) selectively in the northern Baltic Sea, but only at a rate which can affect local 

abundances of E. maritima (Viitasalo and Viitasalo 2004).

Physical

It is possible that Eubosmina maritima is transported over land by recreational boaters. Though it 

is not as likely to foul gear and attach to equipment as the more commonly known Spiny 

Waterflea (Bythotrephes longimanus) and Fishhook Waterflea (Cercopagis pengoi), the same 

responsible maintenance and cleaning methods are recommended to prevent spread between 

water bodies, including cleaning all aquatic equipment with high pressure water (>250 psi) or hot 

water (>50°C) after each use (OFAH n.d.). Electron beam irradiation and ultraviolet light 

treatments have been used to control Spiny and Fishhook Waterfleas in aquatic pathways, and 

are likely effective against E. maritima (USACE 2012b). Another possible non-selective 

pathway control is high water turbidity, which may decrease zooplankton (especially 

Cladoceran) abundances due to the negative effects of suspended clay particles on filtering and 

assimilation rates (Suchy and Hann 2007). 

Chemical

Santagata et al. (2008) found that ballast water exchange methods which flush freshwater 

organisms into euhaline seawater are effective against Eubosmina maritima at a minimum of 24 

PSU (practical salinity units) for two hours in a laboratory simulation. However, E. maritima 

ephippia may remain in residual unpumpable sediment in ballast tanks, and Gray et al. (2005) 

found that exposing zooplankton ephippia to open ocean saline water of 32 ppt (parts per 

thousand) did not reduce egg abundances or consistently affect richness of invertebrates hatched 

from exposed eggs. 

Zooplankton biomass generally increases with increasing eutrophication, so reduction of 

excessive nutrient pollution causing abnormal eutrophication could help control E. coregoni

(Gemza 1995).

A.8 Copepods

Argulus japonicus Thiele, 1900

Gizzard Shad (Dorosoma cepedianum) is a host fish (Poly, 1998) as well as Yellow Perch (Perca 

flavescens), Koi Carp (Cyprinus carpio), and Goldfish (Carassius auratus) (Lesko et al. 2003).

Cyclops strenuus Fischer, 1851

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
158

Control

Biological

Treasurer (1992) found that Eurasian perch (Perca fluviatilis) larvae selectively prey on Cyclops 

strenuus abyssorum in the Scottish lochs Kinord and Davan. The total zooplankton reduction 

observed was minimal, but Treasurer suggested that grazing by larvae is likely to impact 

copepod populations (1992). 

Physical

There are no known physical control methods for this species.

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod resting 

stages. 

Eurytemora affinis Poppe, 1880

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Maes et al. (2005) found that juvenile Herring (Clupea harengus) and Sprat (Sprattus sprattus) 

exhibit top down control of Eurytemora affinis through predation pressure in the Scheldt estuary 

in Belgium. 

Physical

There are no known physical control methods for this species.

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod ephippia. 

Lindley et al. (1999) found that exposure to the organochlorine compounds pentachlorophenol 

(PCP) and 1, 2-dichlorobenzene (DCB), (both common industrial pollutants) accumulated in 

sediment significantly reduced hatching success and nauplii viability of E. affinis eggs. In a study 

of the effect of salinity on toxicity of cadmium to Chesapeake Bay organisms, Hall et al. (1995) 

found that E. affinis is very sensitive to cadmium compared to other estuarine aquatic biota. The 

study documents 96 h LC50 (lethal concentration to 50% of organisms tested) values of 51.6, 

213.2, and 82.9 μg L-1at 5, 15, and 25 ppt (parts per thousand) salinities, respectively (Hall et al. 

1995). Sullivan et al. observed a 96 h LC50 of >120 μg L-1for cadmium and ~30 μg L-1for 

copper on E. affinis at 10 ppt salinity (1983). Sullivan et al. (1983) also noted that reduced 
159

growth rates of E. affinis occur at Cu and Cd doses below the 96 h LC50, which has been 

documented as extending generation length and eventually reducing population size in previous 

studies. The negative effects of the insecticide diflubenzuron (Dimilin®) on E. affinis nauplii 

were documented by Savitz and Wright, who noted a 48 h LC50 of 2.2 μg L-1. The insecticide is 

approved for use against the gypsy moth (Lymantria dispar) and other insect pests by the United 

States Environmental Protection Agency (USEPA), and enters E. affinis habitat through runoff or 

by direct spraying (Savitz and Wright 1994). Diflubenzuron specifically targets the arthropod 

molting process, so the most explicit effects are expected in sub-adult crustaceans. E. affinis was 

lethally affected at levels as low as 0.78 μg L-1(Savitz and Wright 1994). 

Heteropsyllus nr. nunni Coull

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod resting 

stages. 

Megacyclops viridis Jurine, 1820

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Megacyclops viridis is an important prey item for introduced Ruffe (Gymnocephalus cernuus), 

but the Ruffe feeds on a wide variety of benthic organisms, so its feasibility as a biocontrol is 

unknown (Ogle et al. 1995). Yellow Perch (Perca flavescens) and trout perch (Percopsis 

omiscomaycus) are potential predators of M. viridis (Ogle et al. 1995). Hansen and Jeppesen 

(1992) found that a 50% reduction of planktivorous fish biomass (roach, Rutilus rutilus, and 

bream, Abramis brama) affected cyclopoid copepod population directly through reduction in fish 

predation pressure and changes in biological structure of Lake Væng, Denmark. 
160

Physical

There are no known physical control methods for this species. M. viridis is tolerant to salinity of 

a wide range, up to 7.9 g L-1in one study (Wolfram et al. 1999). 

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod ephippia. 

Neoergasilus japonicus Harada, 1930

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

Electron beam irradiation can be used to control microorganisms in aquatic pathways, including 

Neoergasilus japonicus (USACE 2012b). Electron beam irradiation is a non-selective control 

method which exposes water to low doses of radiation using gamma-sterilizers or electron 

accelerators, breaking down DNA in living organisms while leaving behind no by-products 

(USACE 2012b). Ultraviolet (UV) light can also effectively control microorganisms including N. 

japonicus in water treatment facilities and narrow channels, where UV filters can be used to emit 

UV light into passing water, penetrating cell walls and rearranging DNA of microorganisms 

(USACE 2012b).

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod resting 

stages. 

Nitokra hibernica Brady, 1880

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Ballast exchange regulations for international vessels entering the Great Lakes are highly 

efficient at preventing introduction of international nonindigenous species (NIS), but Briski et al. 

have noted that a lack of ballast regulations in domestic shipping within the Great Lakes poses a 

threat to further spread of NIS between lakes (2012). Specifically, they sampled Nitokra

hibernica from ballast water destined for discharge in Lake Superior, where it has not yet been 
161

documented. They suggest domestic regulations be based on ecological boundaries rather than 

geographic and political borders to help prevent spread of NIS within the Great Lakes (Briski et 

al. 2012). 

Control

Biological

Nitokra hibernica has been found in the stomach of Slimy Sculpin (Cottus cognatus) at Yankee 

Reef, Lake Huron, and it is known to be consumed by Rainbow Smelt (Osmerus mordax) in 

Lake Huron, but the significance of this predation on biological control is unknown (Lesko et al. 

2003). 

Physical

There are no known physical control methods for this species. 

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against copepod resting 

stages. 

Nitokra incerta Richard, 1893

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species

Physical

There are no known physical control methods for this species. 

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against resting stages. 

Salmincola lotae Olsson, 1877

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
162

Control

Biological

There are no known biological control methods for this species

Physical

There are no known physical control methods for this species.

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against resting stages. 

Schizopera borutzkyi Monchenko, 1967

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

Schizopera borutzkyi is found in shallow muds and sands at a temperature of 21°C and pH 7.6 in 

its native habitat of the Black Sea Danube River delta, and can tolerate a wide variety of 

salinities (0.04-6%), but there is no research available at this point on the significance of these 

parameters to control (Horvath et al. 2001). 

Electron beam irradiation can be used to control microorganisms in aquatic pathways, including 

S. borutzkyi (USACE 2012b). Electron beam irradiation is a non-selective control method which 

exposes water to low doses of radiation using gamma-sterilizers or electron accelerators, 

breaking down DNA in living organisms while leaving behind no by-products (USACE 2012b). 

Ultraviolet (UV) light can also effectively control microorganisms, including S. borutzkyi, in 

water treatment facilities and narrow channels, where UV filters can be used to emit UV light 

into passing water, penetrating cell walls and rearranging DNA of microorganisms (USACE 

2012b).

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against resting stages. 
163

Skistodiaptomus pallidus Herrick, 1879

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species. 

Chemical

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of copepods. It should be noted that 

the effectiveness of these methods is likely significantly diminished against resting stages. 

A.9 Mysids

Hemimysis anomala G.O. Sars, 1907

Regulations (pertaining to the Great Lakes region)

In Wisconsin, Hemimysis anomala is a prohibited invasive species (WI Administrative Code § 

NR 40.04), meaning that it is unlawful to transport, possess, transfer, or introduce the species 

within or into the state without a permit as defined under Wisconsin Administrative Code § NR 

40.06.

Control

Biological

In Lake Ontario, H. anomala has been documented in the stomachs of Alewife (Alosa 

pseudoharengus), Rock Bass (Ambloplites rupestris), Yellow Perch (Perca flavescens), and 

White Perch (Morone americana) (Brooking et al. 2010, Lantry et al. 2010). Alewives were the 

only predators exhibiting significant consumption of H. anomala, likely due to their nocturnal 

feeding habits (H. anomala exhibits diel vertical migration, remaining near the lakebed during 

the day and emerging at night), and their prior experience consuming the Great Lakes native 

mysid, Mysis relicta, which exhibits similar swimming behavior to H. anomala (Lantry et al. 

2010). Lantry et al. (2010) suggest that as the density and spatial distribution of H. anomala

expands, more Great Lakes fishes will become successful predators. Round Goby (Apollonia 

melanostoma) are accomplished molluscivores, but have also demonstrated specific predatory 

behavior enabling them to consume M. relicta, and may become significant predators of H. 

anomala in the absence of dreissenids (Lantry et al. 2010). H. anomala has also been 

documented in Yellow Perch and White Perch diets in Lake Oneida, New York (Brooking et al. 

2010). The significance of fish predation on control of H. anomala is currently unknown, but 

there is potential for adaptation towards consumption among Great Lakes zooplanktivores, 

especially because it is considered a high-energy food source (Borcherding et al. 2006).
164

Physical

H. anomala exhibits mortality at temperatures below 0° C (Borcherding et al. 2006). Because H. 

anomala is a new, quickly spreading invasive species, preventative measures are encouraged for 

boaters traveling between water bodies, including visually inspecting boats, trailers, and 

equipment for plants, animals, and mud after each use, draining water from the motor, live well, 

bilge, and transom wells while on land, and rinsing all equipment with high pressure (>250 psi) 

or hot (>50°C) water (OFAH n.d.).

Electron beam irradiation can be used to control microorganisms in aquatic pathways, including 

H. anomala (USACE 2012b). Electron beam irradiation is a non-selective control method which 

exposes water to low doses of radiation using gamma-sterilizers or electron accelerators, 

breaking down DNA in living organisms while leaving behind no by-products (USACE 2012b). 

Ultraviolet (UV) light can also effectively control microorganisms including H. anomala in 

water treatment facilities and narrow channels, where UV filters can be used to emit UV light 

into passing water, penetrating cell walls and rearranging DNA of microorganisms (USACE 

2012b).

Chemical

H. anomala tolerates salinity of 0-19 ppt (parts per thousand) (Bij de Vaate et al. 2002). Ellis and 

MacIsaac (2009) tested the salinity tolerance of Great Lakes Invaders in ballast water exchange 

(BWE) simulations. They documented 100% mortality for H. anomala after five hours in a 

simultaneous BWE treatment, in which salinity was gradually increased from 4-30 ppt, and 

100% mortality after three hours in a sequential BWE treatment, in which species are 

immediately exposed to 30 ppt salinity.

The Great Lakes and Mississippi River Interbasin Study (USACE 2012b) suggests that alteration 

of water quality using carbon dioxide, ozone, nitrogen, and/or sodium thiosulfate could be 

effective in preventing upstream and downstream movement of crustaceans.

A.10 Fishes

Alosa aestivalis Mitchill, 1814

Regulations (pertaining to the Great Lakes region)

New York, under federal law, is required to follow the Interstate Fishery Management Plan for 

Shad and River Herring established by the Atlantic States Marine Fisheries Commission 

(Enacted May 2009; ASMFC) and, therefore, are forced to close any non-sustainable commercial 

and recreational fisheries by January 1, 2012 until the Department can prove New York fisheries 

are self-sustaining (NYSDEC 2012b). Management plans are still in review and final 

amendments for each New York fisheries is unknown. New York restricts the use of anadromous 

river Herring (including Blueback) as bait in most waters (6 NYCRR § Part 19). While not listed 

by name, in Ohio it is illegal for any person to possess, import or sell exotic species of fish 

(including Alosa aestivalis) or hybrids thereof for introduction or to release into any body of 

water that is connected to or otherwise drains into a flowing stream or other body of water that 

would allow egress of the fish into public waters, or waters of the state, without first having 

obtained permission (Ohio Administrative Code § 1501:31-19). Ontario takes a whitelist 
165

approach to bait under which all unlisted species (including Alosa aestivalis) are prohibited to 

use as bait (Ontario Fishing Guide 2014).

Control

Most management research relating to Alosa aestivalis appears focused on maintaining 

populations within the native range. Little species-specific information is available on how to 

control this species where it is invasive.

Biological 

There are no known biological control methods for this species. Top-down control by salmonids 

is effective for the related species Alewife, Alosa pseudoharengus.

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Alosa 

aestivalis (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly.

Alosa pseudoharengus Wilson, 1811

Regulations (pertaining to the Great Lakes region)

Alosa pseudoharengus is a regulated invasive species in Minnesota (MN Administrative Rules §

6216.0260). New York restricts the use of Alewife as bait in most waters (6 NYCRR § Part 19). 

While not listed by name, in Ohio it is illegal for any person to possess, import or sell exotic 

species of fish (including Alosa pseudoharengus) or hybrids thereof for introduction or to release 

into any body of water that is connected to or otherwise drains into a flowing stream or other 

body of water that would allow egress of the fish into public waters, or waters of the state, 

without first having obtained permission (Ohio Administrative Code Chapter 1501:31-19). 
166

Ontario takes a whitelist approach to bait under which all unlisted species (including Alosa 

pseudoharengus) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological 

The management response to Great Lakes Alewife overabundance and recurring die-offs was to 

invest in Sea Lamprey (Petromyzon marinus) control and planting of hatchery-reared Pacific 

salmonids (Oncorhynchus spp.) to re-establish top open-water predators (Kocik and Jones 1999,

Hansen and Holey 2002). Older and larger fish tend to be most heavily affected by piscivores, 

while smaller and younger fish remain abundant (Hewett and Stewart 1989). Alewives are now 

managed in part to support the valuable salmonid fishery.

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly.

Apeltes quadracus Mitchill, 1815

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. Ontario takes a whitelist approach to bait under 

which all unlisted species (including Apeltes quadracus) are prohibited to use as bait (Ontario 

Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species.
167

Physical

Netting can be used method of control for some systems, though small size of these fish might 

limit the viability of this option. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Apeltes 

quadracus (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fishes with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Boogaard et al. (2003) found 

that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-

nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined. In another study 

on cumulative toxicity, combinations of Bayer 73 (niclosamide) and TFM with contaminants 

common in the Great Lakes (pesticides, heavy metals, industrial organics, phosphorus, and 

sediments) were found to be mostly additive in toxicity to Rainbow Trout, and one combination 

of TFM, Delnav, and malathion was synergistic, with toxicity magnified 7.9 times (Marking and 

Bills 1985). This highlights the need for managers to conduct on-site toxicity testing and to give 

serious consideration to determining the total toxic burden to which organisms may be exposed 

when using chemical treatments (Marking and Bills 1985). Other non-selective alterations of 

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Carassius auratus Linnaeus, 1758

Regulations (pertaining to the Great Lakes region)

Canadian federal law dictates that no person shall use as bait, or possess for use as bait, in any 

province any live or dead Goldfish (Maritime Provinces Fishery Regulations § SOR 93-55). 

Provincial laws in Ontario and Quebec also state that Goldfish are not to be used as bait (Ontario 

Fishery Regulations § SOR/2007-237 and Quebec Fishery Regulations § SOR 90/214). 

In the state of New York, it is illegal to use or sell Goldfish larvae for bait, and Goldfish larvae 

taken in nets operated pursuant to baitfishing are to be destroyed immediately (NY 

Environmental Conservation Law § 11-1315). In Minnesota, Goldfish are a regulated invasive 

species, making introduction of the species without a permit illegal (MN Administrative Rules § 
168

6216.0260, MN Administrative Rules § 6216.0265). In the state of Pennsylvania, it is unlawful 

to use or possess Goldfish as baitfish while fishing (58 PA Code § 63.44). In the state of 

Wisconsin, Goldfish are a restricted invasive species (WI Administrative Code § NR 40.05). 

Control

Biological

There are no known biological control methods for this species.

Physical

Yamamoto et al. (2006) noted that physical drawdown of water levels has significant negative 

effects on cyprinid spawning abilities in Lake Biwa, Japan. Carassius spp. and Cyprinus carpio

eggs were notably reduced after collection when water levels were lowered by 30 cm, and as 

little as a 10 cm reduction can significantly reduce available shallow, litter-accumulated 

spawning areas preferred by cyprinids (Yamamoto et al. 2006). Potential impacts to native 

species may be significant. This species may be managed by removal (Morgan et al. 2005), 

although this would have to be an ongoing program.

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides (USACE 2012b). 

Goldfish are somewhat more resistant to antimycin A than other fishes (Lennon and Berger 

1964), thus higher concentrations may be required and potential kill of native fishes should be 

given careful consideration. Potassium permanganate (KMnO4) is effective for detoxifying 

antimycin in laboratory waters of pH 6.5 to 9.5. The half-life for antimycin exposed to 1.0 

mg/liter of KMnO4 ranges from 7 to 11 minutes in the different pH waters at 12 C. The 96-h 

LC50's for potassium permanganate (KMnO4) is 3.60 mg/liter with Goldfish, Carassius auratus, 

in laboratory tests. The toxicity of KMnO4 to fish is greatest in water of lower temperatures, in 

harder water, or in higher pH water (Marking and Bills 1975). Treating with antimycin A 

followed by detoxification with potassium permanganate allows restocking within 2 hours 

(Gilderhus et al. 1981).

Relative to other fish species, Goldfish are highly resistant to rotenone. They were the most 

resistant of 21 species tested by Marking and Bills (1976) and second only to Yellow Bullhead in 

Turner’s study (1959).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fishes with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008). Liming has also been 

used to control Goldfish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. This is especially true for Goldfish, which demonstrate a high 
169

resistance to most chemical controls and tolerance for degraded environmental conditions.

Potential effects on non-target plants and organisms, including macroinvertebrates and other 

fishes, should always be deliberately evaluated and analyzed. The effects of combinations of 

management chemicals and other toxicants, whether intentional or unintentional, should be 

understood prior to chemical treatment. Boogaard et al. (2003) found that the lampricides 3-

trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-nitrosalicylanilide (niclosamide) 

demonstrate additive toxicity when combined. In another study on cumulative toxicity, 

combinations of Bayer 73 (niclosamide) and TFM with contaminants common in the Great 

Lakes (pesticides, heavy metals, industrial organics, phosphorus, and sediments) were found to 

be mostly additive in toxicity to Rainbow Trout, and one combination of TFM, Delnav, and 

malathion was synergistic, with toxicity magnified 7.9 times (Marking and Bills 1985). This 

highlights the need for managers to conduct on-site toxicity testing and to give serious 

consideration to determining the total toxic burden to which organisms may be exposed when 

using chemical treatments (Marking and Bills 1985). Other non-selective alterations of water 

quality, such as reducing dissolved oxygen levels or altering pH, could also have a deleterious 

impact on native fishes, invertebrates, and other fauna or flora, and their potential harmful effects 

should therefore be evaluated thoroughly. Goldfish are particularly tolerant of low oxygen levels 

and most other species would be expectedto die first. 

Cyprinus carpio Linnaeus, 1758

Regulations (pertaining to the Great Lakes region)

Cyprinus carpio are a regulated invasive species in Minnesota, making introduction of the 

species without a permit illegal (MN Administrative Rules § 6216.2060, MN Administrative

Rules § 6216.0265). In Wisconsin, Cyprinus carpio is a “restricted species” under NR 40. 

“Restricted species are also subject to a ban on transport, transfer and introduction, but 

possession is allowed, with the exception of fish and crayfish.” Carp may not be collected or 

used for bait in New York or Pennsylvania (NY Environmental Conservation Law § 11-1315; 58 

PA Code § 63.44). Ontario takes a whitelist approach to bait under which all unlisted species 

(including Cyprinus carpio) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Management and control of Common Carp has been well documented through much of 

North America (Meronek et al. 1996, Wydoski and Wiley 1999) with millions of dollars invested 

on research and control (Pimentel et al. 2000).

Biological 

Northern Pike, Esox Lucius, have been used as a biological tool to control Common Carp

recruitment in the Sandhill lakes in Nebraska (Paukert et al. 2003). Spring Viremia of Carp

(SVC) has been suggested as a control of Common Carp in Australia. However, releasing waterborne viral control agents would likely lead to serious disagreement in the scientific and 

management communities, along with the general public due to potential impacts to nontarget 

species (Koehn et al. 2000). Inducible Fatality Genes (IFG) involve breeding carp with a fatal 

genetic weakness to a trigger substance, such as zinc. The fatal gene technology appears to be a 

potentially viable and long-term strategy for the environmentally benign control of carp (Koehn 

et al. 2000).
170

Physical

Removal projects included mechanical harvest by netting (Ritz 1987, Pinto et al. 2005), water 

level manipulation to disrupt spawning (Summerfelt 1999), and exclusion from spawning habitat 

(Lougheed and Chow-Fraser 2001).

Common Carp display jumping behavior when trying to escape entrapment. The Williams cage 

exploits this behavior by selectively removing the jumping carp from other fishes. Tests of the 

Williams cage in Australia proved to be extremely successful. Over the three year testing, the 

Williams cage successfully separated 88% of adult Common Carp and allowed 99.9% native 

species to pass through. The Williams cage is useful in controlling dispersal and abundance of 

Common Carp.

Barriers including electric, bubble curtain, and sonic have been used to exclude carp from 

industrial cooling intake structures (Koehn et al. 2000). Harvesting is only effective if carp are of 

importance by fisheries and anglers. Even if carp are beneficial for harvest, this method is one of 

the least effective methods available (Linfield 1980, Vacha 1998, Koehn et al. 2000, Wedekind 

et al. 2001, Arlinghaus and Mehner 2003). Other physical control methods include traps and 

water level manipulation. Seasonal movements of Common Carp can be exploited to enhance 

management actions, such as removal or inturupting spawning (Taylor et al. 2012). When 

possible, carp can be excluded from an area and then kept out through sorting of fish, which has 

been done since 1997 at the Cootes Paradise Marsh in Hamilton, Ontario (Lougheed et al. 2004). 

Although labor intensive, this method is effective at keeping carp from returning to the marsh.

Chemical

Rotenone is a widely used non-selective chemical used to eliminate Common Carp from a water 

body (Koehn et al. 2000, Sorensen and Stacey 2004, Fajt and Grizzle 1998). 

Antimycin-impregnated baits have been used to target Common Carp (Rach et al. 1994,

Clearwater et al. 2008). The bait pellets consisted of fish meal, a binding agent, antimycin and 

water. Doses of 10 mg antimycin/g bait caused low (19%) to high (74%) mortalities in fish 

feeding voluntarily on 50 g of the toxic bait in each of three earthen ponds.

In laboratory trials, a combination of pH 6.5 and 642 mg/L NaHCO3 was the most effective 

treatment for Rainbow Trout, Brook Trout, and Common Carp, causing the fish to cease 

locomotion and slowing opercular rate within five minutes (Brooke et al. 1978).

Application of different pheromones such as migratory, alarm, and sex may be useful in the 

integrated management of carp (Sorensen and Stacey 2004).

Enneacanthus gloriosus Holbrook, 1855

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. Ontario takes a whitelist approach to bait under 

which all unlisted species (including Enneacanthus gloriosus) are prohibited to use as bait 

(Ontario Fishing Guide 2014).
171

Control

Biological

There are no known biological control methods specific to this species. Top-down control 

(trophic cascade) may be effective for control as it is for native sunfishes.

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on 

Enneacanthus gloriosus (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fishes with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

Gonzalez and Dunson (1989) found that Enneacanthus gloriosus exposed to pH 4.25 and 4.0 

grew at a lower rate (30-40% less growth) than fish of the same species exposed to higher pH 

(4.5, 5.0, 5.8). Survival of E. gloriosus was reduced at pH 3.5 and eliminated at pH 3.25, and the 

authors concluded that E. gloriosus would be excluded from habitats that regularly drop below 

pH 4.0 (Gonzalez and Dunson 1989). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Boogaard et al. (2003) found 

that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-

nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined. In another study 

on cumulative toxicity, combinations of Bayer 73 (niclosamide) and TFM with contaminants 

common in the Great Lakes (pesticides, heavy metals, industrial organics, phosphorus, and 

sediments) were found to be mostly additive in toxicity to Rainbow Trout, and one combination 

of TFM, Delnav, and malathion was synergistic, with toxicity magnified 7.9 times (Marking and 

Bills 1985). This highlights the need for managers to conduct on-site toxicity testing and to give 

serious consideration to determining the total toxic burden to which organisms may be exposed 

when using chemical treatments (Marking and Bills 1985). Other non-selective alterations of 

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 
172

Esox niger Lesueur, 1818

Regulations (pertaining to the Great Lakes region)

The sale of dead Chain Pickerel is prohibited in Quebec under the Quebec Regulation 

Respecting Aquaculture and the Sale of Fish § RRQ, c C-61.1, r 7. Ontario takes a whitelist 

approach to bait under which all unlisted species (including Esox niger) are prohibited to use as 

bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species.

Physical

The Maine Department of Inland Fisheries and Wildlife cites removal of fishing bag limits for 

Chain Pickerel as a management measure in certain lakes, noting that the rationale is biologically 

sound but negligibly effective because few anglers take advantage of the law and capture more 

than 10 pickerel per day (Brokaw 2008). 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides (USACE 2012b).

Esox niger exhibited partial mortality in trials ranging from 0.4-12.1ppb antimycin (Lennon and 

Berger 1970).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fishes with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Boogaard et al. (2003) found 

that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-

nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined. In another study 

on cumulative toxicity, combinations of Bayer 73 (niclosamide) and TFM with contaminants 

common in the Great Lakes (pesticides, heavy metals, industrial organics, phosphorus, and 

sediments) were found to be mostly additive in toxicity to Rainbow Trout, and one combination 

of TFM, Delnav, and malathion was synergistic, with toxicity magnified 7.9 times (Marking and 

Bills 1985). This highlights the need for managers to conduct on-site toxicity testing and to give 

serious consideration to determining the total toxic burden to which organisms may be exposed 

when using chemical treatments (Marking and Bills 1985). Other non-selective alterations of 
173

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Gambusia affinis Baird and Girard, 1853

Regulations (pertaining to the Great Lakes region)

The Western Mosquitofish is prohibited in Wisconsin under Wisconsin Administrative Code § 

NR 40.04, meaning that no person may transport, possess, transfer, or introduce the species 

without authorization. Ontario takes a whitelist approach to bait under which all unlisted species 

(including Gambusia affinis) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Gambusia 

affinis (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly.

Gymnocephalus cernua Linnaeus, 1758

Regulations (pertaining to the Great Lakes region)

Aquarium fish-keeping, production, keeping in captivity, breeding, stocking, transport, sale, or 

purchase of live Ruffe is prohibited in Quebec under the Quebec Regulation Respecting 

Aquaculture and the Sale of Fish § RRQ, c C-61.1, r 7. In Ontario, Ruffe is an invasive fish 
174

under Ontario Fishery Regulations § SOR/2007-237, and, therefore, may not be possessed 

without a license and shall not be used or possessed for use as baitfish. 

In Pennsylvania, it is unlawful to possess live Ruffe or to import or introduce live Ruffe to 

Pennsylvania waters under 58 PA Code § 71.6. It is unlawful to sell, purchase, offer for sale or 

barter for live Ruffe under 58 PA Code § 63.46. Ruffe may not be transported from another state, 

province, or country into Pennsylvania, liberated into a Pennsylvania watershed, or transferred 

between Pennsylvania waters without written permission from the Pennsylvania Fish and Boat 

Commission under 58 PA Code § 73.1. In Ohio, it is unlawful to possess, import, or sell live 

individuals of Ruffe except for research, education, or public display when authorized (Ohio 

Administrative Code § 1501:31-19-01). In Michigan, Ruffe is a prohibited species under 

Michigan NREPA 451 § 324.41301. No person shall knowingly possess a live prohibited 

organism in Michigan except for education, research, or identification purposes as listed in 

Michigan NREPA 451 § 324.41303. It is also unlawful to introduce prohibited organisms in 

Michigan under Michigan NREPA 451 § 324.41305. In Michigan, a violation involving a 

prohibited species is a felony, and a knowing introduction violation with intent to harm is 

punishable with up to five years imprisonment and a $2,000 to $1,000,000 fine (MI NREPA § 

324.41309). In Indiana, Ruffe is classified as an exotic fish under 312 Indiana Administrative 

Code 9-6-7, meaning except as otherwise provided, no individual can import, possess, propagate, 

buy, sell, barter, trade, transfer, loan, or release into public or private waters live fish, recently 

hatched juveniles, viable eggs, or genetic material. In Illinois, Ruffe is listed as an injurious 

species under Illinois Administrative Code 17 § 805.20. It is unlawful to possess, propagate, buy, 

sell, barter, or offer to be bought, sold, bartered, transported, traded, transferred, or loaned an 

injurious species to any person or institution unless a permit is obtained from the Illinois DNR 

(IL Administrative Code 17 § 805.30). In Wisconsin, Ruffe is a restricted species as an 

established non-native, and therefore cannot be transported, possessed, transferred, or introduced 

without a permit (WI Administrative Code § NR 40.05). In Minnesota, Ruffe is a prohibited 

invasive species, meaning it is unlawful (a misdemeanor) to possess, import, purchase, transport, 

or introduce an organism except under permit for control, research, or education (MN

Administrative Rules § 6216.0250). 

Control

Biological

Minnesota and Wisconsin, with advice from the United States Fish and Wildlife Service, 

implemented a top-down control program for Ruffe in the St. Louis River, western Lake 

Superior, in 1989, using Northern Pike (Esox lucius), Walleye (Sander vitreus), Smallmouth 

Bass (Micropterus dolomieui), Brown Bullhead (Ameiurus nebulosus), and Yellow Perch (Perca 

flavescens) (Mayo et al. 1998). A bioenergetics modeling evaluation of the top-down control 

program revealed that although predators ate as much as 47% of Ruffe biomass in one year, they 

avoided Ruffe and were selective for native prey, and were thus unable to halt the increase in 

Ruffe abundance (Mayo et al. 1998). However, the authors noted that Northern Pike and Walleye 

appeared to have potential for top-down control of Ruffe due to a combination of their diets and 

population sizes, and due to indications that they may learn to prey more selectively on Ruffe

(Mayo et al. 1998). 
175

As Mayo et al. (1998) noted, caution is advised when considering top-down biological control as 

a management tool because the stability properties of a system do not just depend on predation, 

but also on the life histories of component species and their interactions.

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but no studies have been found of their effects on 

Oncorhynchus nerka (USACE 2012b).

Evaluation of the effects of common piscicides on Ruffe revealed that the lampricide 3-

trifluoromethyl-4-nitrophenol (TFM) has potential for selective control of the species (Boogaard 

et al. 1996). Ruffe was three to six times more sensitive to TFM than both Yellow Perch (Perca 

flavescens) and Brown Trout (Salmo trutta) (Boogaard et al. 1996). Toxicity tests in May and 

August 1992 on the Brule River, Wisconsin revealed a 12h LC99.9 (concentration at which 

99.9% of organisms are killed after 12 hours) of 5.9 mg/L at normal pH levels (~8.4) and 2.80 

mg/L at low pH levels (Boogaard et al. 1996). Furthermore, at low pH levels (7.7-7.9) 12h 

LC25’s of 7.2 mg/L and 4.6 mg/L were recorded for Yellow Perch and Brown Trout, 

respectively, but at normal pH levels no Brown Trout or Yellow Perch mortality was recorded at 

the highest tested concentration of 8.8 mg/L (Boogaard et al. 1996). A cost benefit analysis of a 

United States Ruffe control program supported TFM as a promising chemical control (Leigh 

1998). However, Dawson et al. (1998) suggest that TFM may have more application for treating 

entire bodies of water rather than localized areas because it tended to repel Ruffe in preference 

tests, allowing them to move to untreated areas. Bottom-release formulations of bayluscide and 

antimycin showed promise for effectiveness in treating localized concentrations of Ruffe, but 

more field testing is needed (Dawson et al. 1998). 

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Boogaard et al. (2003) found 

that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-

nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined. In another study 

on cumulative toxicity, combinations of Bayer 73 (niclosamide) and TFM with contaminants 

common in the Great Lakes (pesticides, heavy metals, industrial organics, phosphorus, and 

sediments) were found to be mostly additive in toxicity to Rainbow Trout, and one combination 
176

of TFM, Delnav, and malathion was synergistic, with toxicity magnified 7.9 times (Marking and 

Bills 1985). This highlights the need for managers to conduct on-site toxicity testing and to give 

serious consideration to determining the total toxic burden to which organisms may be exposed 

when using chemical treatments (Marking and Bills 1985). Other non-selective alterations of 

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Ictiobus cyprinellus Valenciennes in Cuvier and Valenciennes, 1844

Ditching and draining for farmlands, which eliminated shallow lakes, may have reduced

Bigmouth Buffalo populations within their native range in the first half of the 20th century. The 

species is not listed as threatened or endangered in any region of its native or introduced 

distribution.

Regulations (pertaining to the Great Lakes region)

Commercial harvests are regulated in many states. Although rarely caught via hook and line, 

state fishing regulations may apply. Ontario takes a whitelist approach to bait under which all 

unlisted species (including Ictiobus cyprinellus) are prohibited to use as bait (Ontario Fishing 

Guide 2014).

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species

Chemical

There are no known chemical control methods specific to this species. General piscicides (such 

as rotenone) may be used for control, but expect significant kill of non-target species.

Lepisosteus platostomus Rafinesque, 1820

Regulations (pertaining to the Great Lakes region)

In Ohio, a class B aquaculture permit is required to engage in propagation, culture, or sale of 

Shortnose Gar, and two levels of escapement prevention are required if cultured in the Lake Erie 

drainage basin (Ohio Administrative Code § 1501:31-39-01). Ontario takes a whitelist approach 

to bait under which all unlisted species (including Lepisosteus platostomus) are prohibited to use 

as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

There are no known physical control methods for this species. 
177

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but no studies have been found of their effects on Lepisosteus 

platostomus (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Lepomis microlophus Günther, 1859

Regulations (pertaining to the Great Lakes region)

The Redear Sunfish is listed as an approved species for aquaculture production in Michigan 

(Aquaculture Development Act, Michigan Compiled Law § 286.875). In Wisconsin, the Redear 

Sunfish is listed as a restricted species under the definition “nonnative fish in the aquaculture 

industry” (WI Administrative Code § NR 40.02, Ontario takes a whitelist approach to bait under 

which all unlisted species (including Lepomis microlophus) are prohibited to use as bait (Ontario 

Fishing Guide 2014). 40.05). 

Control

Biological

There are no known biological control methods for this species. 

Physical

Removal of catch limits for anglers and public information campaigns could aid control of this 

species.

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Lepomis 

microlophus (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 
178

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Misgurnus anguillicaudatus Cantor, 1842

Regulations (pertaining to the Great Lakes region)

In Michigan, Misgurnus anguillicaudatus is a prohibited species under Michigan Natural 

Resources and Environmental Protection Act (NREPA) 451 § 324.41301. No person shall 

knowingly possess a live prohibited organism in Michigan except for educational, research, or 

identification purposes as listed in Michigan NREPA 451 § 324.41303. It is also unlawful to 

introduce a prohibited organism in Michigan under Michigan NREPA 451 § 324.41305. In 

Michigan, a violation involving a prohibited species is a felony, and a knowing introduction 

violation with intent to harm is punishable with up to five years imprisonment and a $2,000 to 

$1,000,000 fine (MI NREPA § 324.41309). In Wisconsin, M. anguillicaudatus is restricted as a 

nonnative viable fish species in the aquarium trade (WI Administrative Code § NR 40.02 and

40.05). Ontario takes a whitelist approach to bait under which all unlisted species (including 

Misgurnus anguillicaudatus) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but no studies have been found of their effects on M. 

anguillicaudatus (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 
179

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

When planning control and management of this species, special attention should be given to the 

high physiological tolerances which place it in the profile of a successful invader. The Oriental 

Weatherfish can survive temperatures that range from 0-38° C, utilize atmospheric oxygen as a 

facultative air-breather to survive hypoxic conditions, and has been documented surviving 

desiccation for over 81 days with no food, likely perishing from desiccation before starvation 

(Koetsier and Urquhart 2012). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Morone americana Gmelin, 1789

Regulations (pertaining to the Great Lakes region)

Minnesota lists White Perch as a prohibited invasive species (MN Administrative Rules § 

6216.0250). In Ohio it is unlawful for any person to possess, import or sell live White Perch

(Ohio Administrative Code § 1501:31-19). Indiana (312 IN Administrative Code § 9-6-7) 

designates White Perch as an exotic fish - an individual must not import, possess, propagate, 

buy, sell, barter, trade, transfer, loan, or release into public or private waters live fish or recently 

hatched or juvenile live fish or their viable eggs or genetic material. Ontario takes a whitelist 

approach to bait under which all unlisted species (including Morone americana) are prohibited to 

use as bait (Ontario Fishing Guide 2014).

Control

For most waters, the only management recommendation for White Perch is unlimited harvest 

(Smith et al. 2002).

Biological 

Bottom-up control (reduction in food supply) of White Perch usually results in stunting 

accompanied by an increase in population so that the population consists of many small fish 

(Smith et al. 2002).

Physical

There are no known physical control methods for this species.

Chemical

The IJC (2011) recommends rotenone for control of White Perch in rapid response scenarios.
180

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Morone 

americana (USACE 2012b).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five

minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly.

Neogobius melanostomus Pallas, 1814

Regulations (pertaining to the Great Lakes region)

In Ohio (Ohio Administrative Code § 1501:31-19) is is unlawful for any person to possess, 

import or sell live Round Gobies. Michigan (MI NREPA 451, Section 324.41301) and 

Minnesota (MN Administrative Rules § 6216.0250) list Round Goby as a prohibited species. 

Illinois (IL Administrative Code § 17: 805.20) lists Round Gobies as an injurious speciesIn 

Pennsylvania (58 PA Code §71.6), it is illegal to possess, import or introduce round gobies. New

York (6 NYCRR Part 10, Paragraph 10.1(c)(3)) prohibits the use of round gobies as bait. Indiana 

(312 IAC 9-6-7) lists round goby as an exotic fish - an individual must not import, possess, 

propagate, buy, sell, barter, trade, transfer, loan, or release into public or private waters any 

exotic fish (including recently hatched or juvenile live fish or their viable eggs or genetic 

material). Ontario takes a whitelist approach to bait under which all unlisted species (including 

Neogobius melanostomus) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological 

Although many other species consume round goby, no effective and species-specific biocontrol 

has been identified. Among other species, native burbot are being investigated for their potential 

to control goby populations (Madenjian et al 2011).

Physical

Electrical barriers may be successful at limiting the movement of round gobies. In tank studies, 

round gobies did not move through such a barrier (Savino et al 2001).

Chemical
181

The IJC (2011) recommends rotenone for control of round goby in rapid response scenarios.

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered “general” piscicides, but no studies have been found of their effects on Neogobius 

semilunaris (GLMRIS 2012).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the US, not for use 

as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for 5 min. is sufficient to 

anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly.

Other

Rollo et al (2007) reported round gobies will approach a speaker emitting conspecific male calls 

in the field, and female round gobies showed significant attractions to speakers emitting 

conspecific male calls in the laboratory. Therefore round goby phonotaxis could be used to lure 

gravid females to traps. As round gobies will spawn multiple times throughout late spring and 

summer, they should remain receptive to male calls and bioacoustic capture for the entire 

breeding season. 

Notropis buchanani Meek, 1896

Regulations

In Pennsylvania, the ghost shiner is an endangered species. The catching, taking, killing, 

possessing, importing to or exporting from the Commonwealth of Pennsylvania, selling or 

offering for sale or purchasing of any individual of an endangered species, alive or dead, or any 

part thereof, without a special permit, is prohibited. Ontario takes a whitelist approach to bait 

under which all unlisted species (including Notropis buchanani) are prohibited to use as bait 

(Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 
182

Physical

There are no known physical control methods for this species. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but no studies have been found of their effects on this species

(GLMRIS 2012).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the US, not for use 

as euthanasia, and exposure to NaHCO3 concentration of 142-642 mg/L for five minutes is 

sufficient to anaesthetize most fish (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Oncorhynchus gorbuscha Walbaum, 1792

Unlike other Pacific salmon, Pink Salmon were not deliberately stocked for Alewife biocontrol, 

nor are they deliberate stocked. Nonetheless, Pink Salmon descendants of an accidental release 

have become a valued part of the Great Lakes recreational fishery and they are managed 

alongside the other Pacific salmonids. Therefore, Pink Salmon management objectives are not 

geared towards the removal or eradication of the species like with most invaders. 

Regulations (pertaining to the Great Lakes region)

Direct Regulations:

Great Lakes states and provinces have their own specific fishing regulations. Generally, the 

overall goals and objectives of Pacific salmon fishing regulations are the same throughout the 

region i.e., to maintain or enhance a healthy and sustainable salmonid fisheries. Pacific salmon

fishing regulations include daily and season bag limits, size limits, permitted baitfish, manner of 

taking i.e., snagging or hook and line, and designated season dates (See New York DEC, 

Pennsylvania F&BC, Ohio DNR, Michigan DNR, Indiana DNR, Illinois DNR, Minnesota DNR, 

Wisconsin DNR, Ontario MNR, and Quebec MRNF websites for specific fishing regulations). 

Ontario takes a whitelist approach to bait under which all unlisted species (including 

Oncorhynchus gorbuscha) are prohibited to use as bait (Ontario Fishing Guide 2014).
183

Indirect Regulations:

Typically, Pacific salmon regulations are not species specific, but rather regulate the salmonid

fisheries as a whole. Indirect Pacific salmon regulations include mandated salmonid pathogen 

screening tests and baitfish regulations.

Mandatory salmonid pathogen screening tests are implemented in all Great Lakes states and 

provinces. The importation, exportation, and transportation of Pacific salmon is highly regulated 

to control the spread of infectious diseases and parasites such as VHS, Bacterial Kidney Disease

(BKD), and whirling disease (See USGS nonindigenous diseases and parasites factsheets for 

state and provincial regulations).

State and provincial baitfish regulations have aided in preventing the spread of infectious 

disease. Specific and or stricter regulations are placed on baitfish species that are known carriers 

of salmonid pathogens. 

Control

Biological 

Like other Pacific salmon, Pink Salmon prey heavily upon two non-native species in the Great 

Lakes, the Alewife (Alosa pseudoharengus) and Rainbow Smelt (Osmerus mordax). Alewives 

remain a key food source and crucial to the survival of Pacific salmon. Over the past several 

decades, Pacific salmon populations have fluctuated with fluctuating Alewife populations. 

Managing one species significantly impacts the other. Pacific salmon and Alewives have 

significant environmental, socio-economic, and beneficial effects in the Great Lakes and 

therefore integrated management is essential. Rainbow Smelt are also a major component of 

Pacific salmon diet. Similar to Alewives, Pacific salmon and Rainbow Smelt management should 

be integrated. Rainbow Smelt have a high environmental impact and high beneficial effect in the 

Great Lakes. The presence or absence of this species significantly alters predator-prey 

relationships and competition between native species. Managers can also attempt to increase less 

harmful native prey species stocks while allowing harmful invasive prey species to decrease. 

Implementation of this bio-control has potential significant beneficial effects in the Great Lakes 

with few negative impacts (See USGS fact sheets on Alewife and Rainbow Smelt).

Of the 23 nonindigenous diseases and parasites in the Great Lakes, Aeromonas salmonicida, 

Renibacterium salmoninarum, Myxobolus cerebralis, and Novirhabdovirus sp. infections have 

been realized in Great Lakes Pacific salmon, while Heterosporosis sp. and Piscirickettsia cf.

salmonis infections have been realized clinically or outside the Great Lakes. Glugea hertwigi, a 

microsporidian, is known to cause mortality in Rainbow Smelt. Therefore, Pacific salmon

management must include the management of the above pathogens and parasites (See USGS 

factsheets on Aeromonas salmonicida, Renibacterium salmoninarum, Myxobolus cerebralis, 

Novirhabdovirus sp., Heterosporosis sp., Piscirickettsia cf. salmonis, and Glugea hertwigi for 

information on Great Lakes impacts and management).

Physical

Aquaculture facilities manage wild and cultured Pacific salmon stocks through wild stock 

assessments and other methods. Managers are then able to make informed decisions on stocking 

strategies. Research, pathogen screening, and pathogen treatment, etc. is conducted in 

aquaculture facilities (See New York DEC, Pennsylvania F&BC, Ohio DNR, Michigan DNR, 
184

Indiana DNR, Illinois DNR, Minnesota DNR, Wisconsin DNR, Ontario MNR, and Quebec 

MRNF websites for information on salmonid aquaculture and state hatcheries).

Chemical

Chemical controls for Pacific salmon are not intended to eradicate or kill the species but rather to 

protect it against infectious disease. Typically, depending on the target species, chemicals 

controls are only effective in aquaculture or similar systems. Examples of chemicals used and 

include Furogen®, chlorination, and disinfectants. 

Oncorhynchus kisutch Walbaum, 1792

Pacific salmon were first introduced to the Great Lakes in the 1960s to manage Alewife

populations. Soon after, the multi-million dollar Great Lakes Pacific salmon sportfishery was 

established and is now one of the largest economic sectors in the region. Therefore, Pacific 

salmon management objectives are not geared towards the removal or eradication of the species 

like with most invaders, but rather to maintain or enhance the health and stability of the fisheries. 

Managers and citizens understand that with over 180 nonindigenous species, the Great Lakes are 

not the same ecosystem they once were. Management efforts still focus on the prevention and 

eradication of harmful invaders, but also realize that non-native Pacific salmon fisheries are one 

of the driving economic forces in the Great Lakes and managers need to account for this. Pacific 

salmon management is extremely diverse, integrated, and cascading and is therefore these are the 

most heavily regulated species (direct and indirectly) in the Great Lakes.

Regulations (pertaining to the Great Lakes region)

Direct Regulations:

Great Lakes states and provinces have their own specific fishing regulations. Generally, the 

overall goals and objectives of Pacific salmon fishing regulations are the same throughout the 

region i.e., to maintain or enhance a healthy and sustainable salmonid fisheries. Pacific salmon

fishing regulations include daily and season bag limits, size limits, permitted baitfish, manner of 

taking i.e., snagging or hook and line, and designated season dates (See New York DEC, 

Pennsylvania F&BC, Ohio DNR, Michigan DNR, Indiana DNR, Illinois DNR, Minnesota DNR, 

Wisconsin DNR, Ontario MNR, and Quebec MRNF websites for specific fishing regulations).

Ontario takes a whitelist approach to bait under which all unlisted species (including 

Oncorhynchus kisutch) are prohibited to use as bait (Ontario Fishing Guide 2014).

Indirect Regulations:

Typically, Pacific salmon regulations are not species specific, but rather regulate the salmonid

fisheries as a whole. Indirect Pacific salmon regulations include mandated salmonid pathogen 

screening tests and baitfish regulations.

Mandatory salmonid pathogen screening tests are implemented in all Great Lakes states and 

provinces. The importation, exportation, and transportation of Pacific salmon is highly regulated 

to control the spread of infectious diseases and parasites such as VHS, Bacterial Kidney Disease

(BKD), and whirling disease (See USGS nonindigenous diseases and parasites factsheets for 

state and provincial regulations).
185

State and provincial baitfish regulations have aided in preventing the spread of infectious 

disease. Specific and or stricter regulations are placed on baitfish species that are known carriers 

of salmonid pathogens. 

Control

Biological 

Pacific salmon prey heavily upon two non-native species in the Great Lakes, the Alewife (Alosa 

pseudoharengus) and Rainbow Smelt (Osmerus mordax). Alewives remain a key food source 

and crucial to the survival of Pacific salmon. Over the past several decades, Pacific salmon

populations have fluctuated with fluctuating Alewife populations. Managing one species 

significantly impacts the other. Pacific salmon and Alewives have significant environmental, 

socio-economic, and beneficial effects in the Great Lakes and therefore integrated management 

is essential. Rainbow Smelt are also a major component of Pacific salmon diet. Similar to 

Alewives, Pacific salmon and Rainbow Smelt management should be integrated. Rainbow Smelt

have a high environmental impact and high beneficial effect in the Great Lakes. The presence or 

absence of this species significantly alters predator-prey relationships and competition between 

native species. Managers can also attempt to increase less harmful native prey species stocks 

while allowing harmful invasive prey species to decrease. Implementation of this bio-control has 

potential significant beneficial effects in the Great Lakes with few negative impacts (See USGS 

fact sheets on Alewife and Rainbow Smelt).

Of the 23 nonindigenous diseases and parasites in the Great Lakes, Aeromonas salmonicida, 

Renibacterium salmoninarum, Myxobolus cerebralis, and Novirhabdovirus sp. infections have 

been realized in Great Lakes Pacific salmon, while Heterosporosis sp. and Piscirickettsia cf.

salmonis infections have been realized clinically or outside the Great Lakes. Glugea hertwigi, a 

microsporidian, is known to cause mortality in Rainbow Smelt. Therefore, Pacific salmon

management must include the management of the above pathogens and parasites (See USGS 

factsheets on Aeromonas salmonicida, Renibacterium salmoninarum, Myxobolus cerebralis, 

Novirhabdovirus sp., Heterosporosis sp., Piscirickettsia cf. salmonis, and Glugea hertwigi for 

information on Great Lakes impacts and management).

Physical

Aquaculture facilities manage wild and cultured Pacific salmon stocks through wild stock 

assessments and other methods. Managers are then able to make informed decisions on stocking 

strategies. Research, pathogen screening, and pathogen treatment, etc. is conducted in 

aquaculture facilities (See New York DEC, Pennsylvania F&BC, Ohio DNR, Michigan DNR, 

Indiana DNR, Illinois DNR, Minnesota DNR, Wisconsin DNR, Ontario MNR, and Quebec 

MRNF websites for information on salmonid aquaculture and state hatcheries).

Chemical

Chemical controls for Pacific salmon are not intended to eradicate or kill the species but rather to 

protect it against infectious disease. Typically, depending on the target species, chemicals 

controls are only effective in aquaculture or similar systems. Examples of chemicals used and 

include Furogen®, chlorination, and disinfectants. 
186

Oncorhynchus mykiss Walbaum, 1792

Regulations (pertaining to the Great Lakes region)

Federal law in Canada regulates Rainbow Trout as a game fish (CRC § c.1120). In Quebec, it is 

illegal to stock Rainbow Trout in certain bodies of water as listed by Quebec RRQ § c C-61.1, r 

7. The sale of dead Rainbow Trout is also prohibited in Quebec (Quebec RRQ § c C-61.1, r 7). 

In Ontario, Rainbow Trout is regulated as an eligible species for aquaculture (Ontario Regulation

§ 664/98). Ontario takes a whitelist approach to bait under which all unlisted species (including 

Oncorhynchus mykiss) are prohibited to use as bait (Ontario Fishing Guide 2014).

In New York, trout, including Rainbow Trout, shall not be bought and sold, excepting cases in 

which a hatchery permit is issued, as described under New York Environmental Conservation 

Law § 11-1909. In Ohio, it is unlawful to take or possess Rainbow Trout less than 12 inches in 

length while on Lake Erie or its tributaries, including all streams in the entire drainage basin, 

excepting Cold Creek upstream of state route two located in Erie county, and Beaver creek in 

Seneca County. It is also unlawful to take or possess Rainbow Trout less than twelve inches in 

length while on the Mad River or its tributaries (Ohio Administrative Code § 1501:31-13-09). In 

Michigan, Rainbow Trout is an approved species for aquaculture production (MI Compiled Law

§ 286.875). In Wisconsin, Rainbow Trout is restricted as a nonnative fish species in the 

aquaculture industry, and therefore cannot be transported, possessed, transferred, or introduced 

without a permit (WI Admin Code § NR 40.05). 

Control

Biological

There are no known biological control methods for this species. 

Physical

In streams and rivers, barriers can be constructed and natural barriers augmented to prevent 

upstream migration of trout and aid management and eradication efforts. In a report on a 

successful trout removal program involving a combination of piscicides (rotenone) for 

eradication and barriers for prevention of re-invasion, Lintermans and Raadik (2003) noted three

key aspects of successful barriers: a 1.5 m or greater vertical drop; direction of water flow 

towards the middle in higher flows with no slower overland flow passing down the banks; and no 

deep pool below the barrier from which trout could jump. Rainbow Trout often rely on spawning 

streams and small tributaries for reproduction, and removal of access to such streams could 

reduce or potentially eliminate populations in downstream bodies of water (Champion et al. 

2002). The United States National Park Service uses physical removal through electrofishing to 

manage Rainbow Trout and Brown Trout populations that threaten native Brook Trout in 

Shenandoah National Park, Virginia (NPS 2011). 

Chemical

Antimycin A (available as Fintrol®) is a registered piscicide in the United States that is 

documented as highly effective against scaled fishes, including Rainbow Trout (Finlayson et al. 

2002). Elimination of trout is achievable in a contact time of two hours at 5 μg/L (5 ppb), or in 

one hour at 10 ppb (Gilderhus 1972, Finlayson et al. 2002). Antimycin is most effective in small 

streams, shallow ponds, and alpine lakes where there is ample mixing and an adequate contact 

time can be achieved (Finlayson et al. 2002, Gilderhus 1972). Antimycin does not seem to repel 

fish like rotenone (another registered piscicide, available as Noxfish®) does, and rapidly breaks 
187

down by hydrolysis in natural waters (Finlayson et al. 2002). Disadvantages of antimycin include 

increasing ineffectiveness in waters with higher pH (>8), streams with significant gradients (80-

150 m elevation drop), and large lakes where good mixing and contact time cannot be 

established (Finlayson et al. 2002). Rotenone is also effective against Rainbow Trout, but at 

much higher concentrations, with 50 μg/L required to eliminate trout in a 2 hour contact time 

(Gilderhus 1972, Finlayson et al. 2002). Antimycin may be preferred because of the lower dose 

required. Antimycin and rotenone are non-selective, and toxicity to other fishes and aquatic 

invertebrates will vary.

Lintermans and Raadik (2003) provide a detailed account of Rainbow Trout elimination 

programs using rotenone in order to protect fishes of the family Galaxiidae, conducted in two 

separate areas of Australia. In 1992, Rainbow Trout were removed from 2.4 km of Lees Creek, 

Australian Capital Territory using a 5% rotenone emulsion at concentrations of approximately 

0.05 parts per million (ppm) (Lintermans and Raadik 2003). The creek was treated in 500 m 

sections, with 300-350 mL of rotenone added over a 15-minute period to each section, and with 

mesh stop nets placed after each section to prevent downstream reinvasion of trout (Lintermans 

and Raadik 2003). An oxidant (350-500 g potassium permanganate) was added to the stream 

when rotenone reached the downstream limit of treatment sections to remove the toxicant 

(Lintermans and Raadik 2003). To prevent trout reinvasion of the treated area, a downstream 

weir was augmented with a heavy steel grill to present a 1.75 m vertical barrier (Lintermans and 

Raadik 2003). Complete eradication was accomplished at the Lees Creek site, and despite heavy 

impacts on aquatic macroinvertebrates, benthic macroinvertebrates remained in significant 

numbers (Lintermans and Raadik 2003). In 1995, a total of 20 km of stream length (seven

different streams) was treated in Victoria, with a total of 60 L rotenone used, neutralized with 

1100 kg of potassium permanganate, and with rotenone volumes ranging from 0.3 to 0.5 L per 

100 m of stream (Lintermans and Raadik 2003). Areas in which trout and galaxiids overlapped 

were not treated with piscicides; they were instead intensively electrofished to more selectively 

remove Rainbow Trout (Lintermans and Raadik 2003). 

The effects of combinations of management chemicals and other toxicants, whether intentional 

or unintentional, should be understood before pursuing chemical treatment options. Boogaard et 

al. (2003) found that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro4’-nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined, with Rainbow 

Trout demonstrating 12h LC50s of 8.40-10.6 mg/L in response to treatments of TFM and 5.00-

5.05 mg/L in response to treatments of a TFM/1% niclosamide combination in lab tests 

(Boogaard et al. 2003). In another study on cumulative toxicity, combinations of Bayer 73 

(niclosamide) and TFM with contaminants common in the Great Lakes (pesticides, heavy metals, 

industrial organics, phosphorus, and sediments) were found to be mostly additive in toxicity to 

Rainbow Trout, and one combination of TFM, Delnav, and malathion was synergistic, with 

toxicity magnified 7.9 times (Marking and Bills 1985). This highlights the need for managers to 

conduct on-site toxicity testing and to give serious consideration to determining the total toxic 

burden to which organisms may be exposed when using chemical treatments (Marking and Bills 

1985).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 
188

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). 

Laboratory trials demonstrated a combination of pH 6.5 and 642 mg/L NaHCO3 was the most 

effective treatment for Rainbow Trout (Clearwater et al. 2008). CO2 is approved only for use as 

an anaesthetic for cold, cool, and warm water fishes in the United States. It is not approved for 

use as euthanasia (Clearwater et al. 2008).

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. Other 

non-selective alterations of water quality, such as reducing dissolved oxygen levels or altering 

pH, could also have a deleterious impact on native fishes, invertebrates, and other fauna or flora, 

and their potential harmful effects should therefore be evaluated thoroughly. 

Oncorhynchus nerka (Walbaum in Artedi, 1792)

Regulations (pertaining to the Great Lakes region)

In Canada, Sockeye Salmon is a game fish as designated by the National Parks of Canada 

Fishing Regulations § CRC, c. 1120. Sockeye Salmon is also a game fish in the provinces of 

Ontario and Quebec (Ontario Fish Regulations § SOR/2007-237; Quebec Fish Regulations § 

SOR/90-214). Ontario takes a whitelist approach to bait under which all unlisted species 

(including Oncorhynchus nerka) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

Sockeye Salmon are a homing species, returning to their native stream for reproduction. Barriers 

can be constructed and/or natural barriers augmented to prevent upstream migration and aid 

management and eradication efforts, though little research exists on effective barriers for 

Oncorhynchus nerka. Lintermans and Raadik (2003) noted three key aspects of successful 

barriers in relation to a Rainbow Trout control program: a 1.5 m or greater vertical drop; 

direction of water flow towards the middle in higher flows with no slower overland flow passing 

down the banks; and no deep pool below the barrier from which fish could jump. 

The United States Army Corps of Engineers Great Lakes and Mississippi River Interbasin Study 

(USACE GLMRIS) notes the potential effectiveness of sensory deterrent systems in providing 

barriers to fish migration or eliciting fish movements (USACE 2012b). Specifically, the success 

of underwater strobe lights as studied by Maiolie et al. (2001) is cited. Testing conducted on 

wild, free-ranging O. nerka in their natural pelagic habitat in two large Idaho lakes revealed that 

fish moved an average of 30-136 m away from lights in waters with secchi transparency of 2.8 to 

17.5 m, with an 80% reduction in fish density within 30 m of the strobe lights (Maiolie et al. 

2001). Many large scale strobe systems consist of four individual lights that flash at a rate of 450

flashes/minute, with an approximate intensity of 2634 lumens/flash (USACE 2012b). Maiolie et 

al. (2001) tested flash rates of 300, 360, and 450 flashes/minute.
189

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but no studies have been found of their effects on O. nerka 

(USACE 2012b).

Exposure to niclosamide, registered in the United States as a granular lampricide, wettable 

powder, technical grade product, and an emulsifiable concentrate, is known to be toxic to all fish 

species at 0.5 mg/L after a 48-h exposure (Clearwater et al. 2008). There are no available studies 

of its specific effects on Sockeye Salmon, though it has been used for control of aquatic snails, 

Zebra Mussels, and oligochaetes, and is also toxic to many crayfish, frogs, clams, algae, and 

other amphibian and fish species (Clearwater et al. 2008).

In a study on removal of toxic chemicals from water using activated carbon, Dawson et al. 

(1976) found that granular activated carbon is saturated by rotenone at 0.1 mg of Noxfish per 

gram carbon, and cited other studies documenting 0.94-1.32 mg of Noxfish adsorbed per gram of 

carbon. Antimycin was efficiently absorbed and did not saturate carbon because of the low doses 

used (Dawson et al. 1976). 

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). 

Salmonids are considered to be among the most sensitive fishes to low dissolved oxygen levels, 

with a DO concentration of 1-3 mg/L sufficient to cause mortality or loss of equilibrium 

(Clearwater et al. 2008). However, CO2 is approved only for use as an anaesthetic for cold, cool, 

and warm water fishes the United States, for use as euthanasia (Clearwater et al. 2008). Exposure 

to NaHCO3 concentration of 142-642 mg/L for five minutes is sufficient to anaesthetize most 

fish (Clearwater et al. 2008). 

Low pH is known to affect fish behavior. Ikuta et al. (2003) documented the effects of low pH on 

Sockeye Salmon, noting that salmon would not swim upstream into areas of pH lower than 6.0. 

Acute exposure to low pH levels can directly kill fish by discharge of sodium and chloride ions 

from body fluid, and sub-lethal levels can affect reproduction (Ikuta et al. 2003). In the case of 

Sockeye Salmon, weak acidic conditions of <pH 6 were enough to depress the prespawning 

behavior of swimming upstream (Ikuta et al. 2003). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target organisms, such as 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. Other 

non-selective alterations of water quality, such as reducing dissolved oxygen levels or altering 

pH, could also have a deleterious impact on native fishes, invertebrates, and other fauna, and 

their potential harmful effects should therefore be evaluated thoroughly. 

Oncorhynchus tshawytscha Walbaum in Artedi, 1792

Pacific salmon were first introduced to the Great Lakes in the 1960s to manage Alewife

populations. Soon after, the multi-million dollar Great Lakes Pacific salmon sportfishery was 
190

established and is now one of the largest economic sectors in the region. Therefore, Pacific 

salmon management objectives are not geared towards the removal or eradication of the species 

like with most invaders, but rather to maintain or enhance the health and stability of the fisheries. 

Managers and citizens understand that with over 180 nonindigenous species, the Great Lakes are 

not the same ecosystem they once were. Management efforts still focus on the prevention and 

eradication of harmful invaders, but also realize that non-native Pacific salmon fisheries are one 

of the driving economic forces in the Great Lakes and managers need to account for this. Pacific 

salmon management is extremely diverse, integrated, and cascading and is therefore these are the 

most heavily regulated species (direct and indirectly) in the Great Lakes.

Regulations (pertaining to the Great Lakes region)

Direct Regulations:

Great Lakes states and provinces have their own specific fishing regulations. Generally, the 

overall goals and objectives of Pacific salmon fishing regulations are the same throughout the 

region i.e., to maintain or enhance a healthy and sustainable salmonid fisheries. Pacific salmon

fishing regulations include daily and season bag limits, size limits, permitted baitfish, manner of 

taking i.e., snagging or hook and line, and designated season dates (See New York DEC, 

Pennsylvania F&BC, Ohio DNR, Michigan DNR, Indiana DNR, Illinois DNR, Minnesota DNR, 

Wisconsin DNR, Ontario MNR, and Quebec MRNF websites for specific fishing regulations).

Ontario takes a whitelist approach to bait under which all unlisted species (including 

Oncorhynchus tshawytscha) are prohibited to use as bait (Ontario Fishing Guide 2014).

Indirect Regulations:

Typically, Pacific salmon regulations are not species specific, but rather regulate the salmonid

fisheries as a whole. Indirect Pacific salmon regulations include mandated salmonid pathogen 

screening tests and baitfish regulations.

Mandatory salmonid pathogen screening tests are implemented in all Great Lakes states and 

provinces. The importation, exportation, and transportation of Pacific salmon is highly regulated 

to control the spread of infectious diseases and parasites such as VHS, bacterial kidney disease 

(BKD), and whirling disease (See USGS nonindigenous diseases and parasites factsheets for 

state and provincial regulations).

State and provincial baitfish regulations have aided in preventing the spread of infectious 

disease. Specific and or stricter regulations are placed on baitfish species that are known carriers 

of salmonid pathogens. 

Control

Biological 

Pacific salmon prey heavily upon two non-native species in the Great Lakes, the Alewife (Alosa 

pseudoharengus) and Rainbow Smelt (Osmerus mordax). Alewives remain a key food source 

and crucial to the survival of Pacific salmon. Over the past several decades, Pacific salmon

populations have fluctuated with fluctuating Alewife populations. Managing one species 

significantly impacts the other. Pacific salmon and Alewives have significant environmental, 

socio-economic, and beneficial effects in the Great Lakes and therefore integrated management 

is essential. Rainbow Smelt are also a major component of Pacific salmon diet. Similar to 

Alewives, Pacific salmon and Rainbow Smelt management should be integrated. Rainbow Smelt

have a high environmental impact and high beneficial effect in the Great Lakes. The presence or 
191

absence of this species significantly alters predator-prey relationships and competition between 

native species. Managers can also attempt to increase less harmful native prey species stocks 

while allowing harmful invasive prey species to decrease. Implementation of this bio-control has 

potential significant beneficial effects in the Great Lakes with few negative impacts (See USGS 

fact sheets on Alewife and Rainbow Smelt).

Of the 23 nonindigenous diseases and parasites in the Great Lakes, Aeromonas salmonicida, 

Renibacterium salmoninarum, Myxobolus cerebralis, and Novirhabdovirus sp. infections have 

been realized in Great Lakes Pacific salmon, while Heterosporosis sp. and Piscirickettsia cf.

salmonis infections have been realized clinically or outside the Great Lakes. Glugea hertwigi, a 

microsporidian, is known to cause mortality in Rainbow Smelt. Therefore, Pacific salmon

management must include the management of the above pathogens and parasites (See USGS 

factsheets on Aeromonas salmonicida, Renibacterium salmoninarum, Myxobolus cerebralis, 

Novirhabdovirus sp., Heterosporosis sp., Piscirickettsia cf. salmonis, and Glugea hertwigi for 

information on Great Lakes impacts and management).

Physical

Aquaculture facilities manage wild and cultured Pacific salmon stocks through wild stock 

assessments and other methods. Managers are then able to make informed decisions on stocking 

strategies. Research, pathogen screening, and pathogen treatment, etc. is conducted in 

aquaculture facilities (See New York DEC, Pennsylvania F&BC, Ohio DNR, Michigan DNR, 

Indiana DNR, Illinois DNR, Minnesota DNR, Wisconsin DNR, Ontario MNR, and Quebec 

MRNF websites for information on salmonid aquaculture and state hatcheries).

Chemical

Chemical controls for Pacific salmon are not intended to eradicate or kill the species but rather to 

protect it against infectious disease. Typically, depending on the target species, chemicals 

controls are only effective in aquaculture or similar systems. Examples of chemicals used and 

include Furogen®, chlorination, and disinfectants. 

Osmerus mordax Mitchill, 1814

Regulations (pertaining to the Great Lakes region)

Use of Rainbow Smelt is regulated in the Canadian province of Quebec under Quebec Fish 

Regulations § SOR/90-214. A population of Rainbow Smelt in an area south of the St. Lawrence 

estuary is designated a vulnerable wildlife species in Quebec under Quebec Statutes and 

Regulations RRQ § c E-12.01, r 2. The sale of dead Rainbow Smelt is prohibited in Quebec by 

Quebec Statutes and Regulations RRQ § c C-61.1, r 7. In Ontario, Rainbow Smelt use as bait and 

non-angling fishing methods are regulated by Canada Federal Statutes and Regulations SOR § 

2007-237. Ontario takes a whitelist approach to bait under which all unlisted species (including 

Osmerus mordax) are prohibited to use as bait (Ontario Fishing Guide 2014).

In the state of New York, it is unlawful to use Rainbow Smelt as bait except as provided in New 

York Codes, Rules, and Regulations 6 § 19.2. Furthermore, it is unlawful to take Rainbow Smelt

for sale as bait or to sell as bait, except as otherwise provided as without a pursuant license as 

defined in New York Environmental Conservation Law § 11-1315. In Pennsylvania, the use of 

commercial trap nets under license to capture Rainbow Smelt is regulated by Pennsylvania

Administrative Code § 69.33. In Ohio, Rainbow Smelt is defined as a commercial fish and an 
192

unrestricted species under Ohio Administrative Code 1501 § 31-1-02. Commercial fish are 

permitted to be taken, possessed, bought, or sold unless otherwise restricted in Ohio code. In 

Indiana, the Rainbow Smelt sport fishing season on Lake Michigan is defined as March 1-May 

30, with capture allowed only by the use of dip nets, seines, or nets with limitations provided in 

312 Indiana Administrative Code § 9-7-2. There is otherwise no bag limit, possession limit, or 

size limit, as defined under 312 Indiana Administrative Code § 9-7-14. In Illinois, the sportfishing season of Rainbow Smelt is defined as March 1-April 30 under Illinois Administrative

Code 17-1 § 810.10. In Wisconsin, Rainbow Smelt is defined as an established non-native fish 

species in Wisconsin Administrative Code § NR 40.02, and is restricted per the above definition 

by Wisconsin Admin Code § NR 40.05. In Minnesota, Rainbow Smelt is a regulated invasive 

species under Minnesota Administrative Rules § 6216.0260. 

Control

Biological

Several species of non-native salmonids have been introduced to the Great Lakes, beginning in 

the 1960s, to control invasive Rainbow Smelt (USACE 2012b). Rainbow Smelt is heavily 

consumed by Atlantic Salmon (Salmo salar), Lake Trout (Salvelinus namaycush), Brook Trout 

(S. fontinalis), Coho Salmon (Oncorhynchus kisutch), Chinook Salmon (O. tshawytcha), 

Rainbow Trout (O. mykiss), Brown Trout (Salmo trutta), Splake (Brook Trout x Lake Trout), 

Burbot (Lota lota), Walleye (Sander vitreus), Northern Pike (Esox lucius), and many other 

freshwater piscivores (Stewart et al. 1981, Brandt and Madon 1986, Crossman 1991, He and 

LaBar 1994, Kirn and LaBar 1996, USACE 2012b). However, the significance of piscivore 

predation on Rainbow Smelt has only been studied for a few species. Observed Atlantic Salmon

predation on smaller Rainbow Smelt, as well as bioenergetics modeling suggesting that by age 

four, cumulative piscivory by Atlantic Salmon was nearly 10-fold greater than that of Lake Trout

of the same age, implies its greater usefulness for management of Rainbow Smelt (Kirn and 

LaBar 1996). While Lake Trout consume large amounts of Rainbow Smelt, almost exclusively 

so in some studies, the species is believed to provide little potential for responsive management 

manipulation outside of stabilizing fluctuating prey populations, due to the long cycle of its 

predatory effect (peaking 3-5 years after stocking, lasting 7-8 years) (Stewart et al. 1981, He and 

LaBar 1994, Kirn and LaBar 1996). Chinook Salmon have been successfully used to eradicate 

Rainbow Smelt from small lakes in New Hampshire in 1936 (Stewart et al. 1981). Because the 

trade-off between fish species as agents of biological control is not directly correlated with 

consumption, management decisions involving shifts between species should not take 

consumption solely into account (Stewart et al. 1981).

Physical

The United States Army Corps of Engineers Great Lakes and Mississippi River Interbasin Study 

notes the potential effectiveness of sensory deterrent systems in providing barriers to fish 

migration or eliciting fish movements (USACE 2012b). In situ testing of two models of strobe 

lights as a deterrent preventing entrainment of Rainbow Smelt through Oahe Dam, Lake Oahe, 

South Dakota demonstrated successful avoidance of 15-21 m horizontally and six meters

vertically by Rainbow Smelt (Hamel et al. 2008). Many large scale strobe systems consist of four 

individual lights that flash at a rate of 450 flashes/minute, with an approximate intensity of 2634 

lumens/flash (USACE 2012b). Hamel et al. (2008) tested the AGL FH-901 flashhead, which 

consists of four horizontal lights positioned at 90 degree angles, flashing 450 times/min at 2,634 

lumens/flash, and the newer AGL FH-920 flashhead, which consists of an omnidirectional 
193

vertical light tube, covering a full 360 degrees with 360 flashes/min at 6,585 lumens. When using 

physical deterrents as barriers, combining methods can increase effectiveness, as was the case for 

Patrick et al. (1985), who found that Rainbow Smelt and other pelagic fishes were successfully 

deterred by a barrier combining air bubbles and strobe lights. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides (USACE 2012b). Marking et al. (1983) found that the three 

most effective registered chemicals for potential use in control of Rainbow Smelt eggs and larvae 

are rotenone, potassium permanganate, and chlorine, respectively. In exposures of 6-24 hours, all 

chemicals were effective at concentrations from 5 to >10 mg/L (Marking et al. 1983). Rotenone 

demonstrated a 96h LC50 of 0.015 mg/L for Rainbow Smelt eggs and 0.001 mg/L for larvae 

(derived calculating only the activity of rotenone in 5% Noxfish solution) (Marking et al. 1983). 

Potassium permanganate demonstrated 96h LC50s of 0.074 mg/L and 0.075 mg/L for eggs and 

larvae, respectively. Chlorine demonstrated 96h LC50s of 0.14 mg/L for eggs and 0.31 mg/L for 

larvae (Marking et al. 1983). Temperature, pH, and hardness of water all affected toxicity of 

rotenone and potassium permanganate, with higher temperatures, softer water, and higher pH 

increasing toxicity (Marking et al. 1983). It should be noted that tests were carried out in a 

laboratory, but natural waters usually contain oxidizable material, which produces a demand on 

chlorine and reduces it to a less active form (Marking et al. 1983).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 

not for use as euthanasia (Clearwater et al. 2008). Exposure to NaHCO3 concentration of 142-

642 mg/L for five minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatmentOther non-selective alterations of 

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Petromyzon marinus (Linnaeus, 1758)

Regulations (pertaining to the Great Lakes region)

In Minnesota, sea lamprey is a prohibited species and therefore it is unlawful to possess, import, 

purchase, transport, or introduce this species except under a permit for disposal, control, 

research, or education (MDNR 2012). In Ohio it is illegal to possess, import or sell live lamprey 

(OAC Chapter 1501:31-19). New York (NY ECL 11-1315, 6a) prohibits the use of lamprey 
194

larvae as bait. Ontario takes a whitelist approach to bait under which all unlisted species 

(including Petromyzon marinus) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Since the bi-national Sea Lamprey Control program (managed by the Great Lakes Fishery 

Commission) was started in the 1950s, populations have been reduced by 90%, and fish survival 

and spawning have increased (Page and Laird 1993, Smith 1985). It is impossible to completely 

eradicate sea lamprey from the Great Lakes; however, continuous control efforts can minimize 

their impacts on the ecosystem and fisheries (FOC 2009).

Biological

An effective bio-control of sea lamprey is the implementation of the sterile-male-release 

program. Male sea lamprey are captured during spawning runs, sterilized using bisazir, and 

released to compete with fertile males for mating; thus reducing egg fertilization. Released 

males are sterilized past their parasitic phase and do not return to the lake. (FOC 2009, GLFHC 

2000a). A potential alternative to bisazir is a lamprey GnRH antagonist. However, further 

research of this alternative sterilizing agent is necessary (Bergstadt and Twohey 2007).

Physical

Barriers and traps have been effective controls of sea lamprey since the 1950s. Barrier options 

include mechanical weirs, electrical barriers, low-head barriers, adjustable crest barriers, and 

velocity barriers (GLFHC 2000, Scott and Crossman 1973, Smith and Tibbles 1980). Traps are 

often used in association with barriers to capture sea lamprey while allowing desired species to 

continue upstream (FOC 2009, GLFHC 2000b). Barriers have reduced the need for lampricide 

applications (GLFC 2012). Once captured, sea lamprey are killed, used for research, or used in 

sterile-male -release programs.

Chemical

Beginning in the late 1950s, sea lampreys began to be successfully controlled by use of the 

lampricide 3-trifluoromethyl-4-nitrophenol (TFM), a chemical agent that kills larval lampreys in 

their stream habitats (Smith and Tibbles 1980). The lampricide has reduced the population by 

over 90% of the 1961 peak (Scott and Crossman 1973). However, continued use of TFM is 

required to keep populations under control (Becker 1983, Scott and Crossman 1973). TFM is 

sometimes harmful to other fish (e.g., walleye) as well as to the larvae of nonparasitic native 

lamprey species (Becker 1983). Bayluscide (niclosamide) treatments in deltas are also a widely 

used and an effective control of sea lamprey larvae (NYSDEC 2012).

Other

To increase the efficacy of lampricide treatments, streams and rivers are frequently assessed for 

larvae density to help determine the application sites (FOC 2009, GLFC 2012).

Note: Check state/provincial and local regulations for the most up-to-date information regarding 

permits for control methods. Follow all label instructions.
195

Phenacobius mirabilis (Girard, 1856)

Regulations

There are no known regulations for this species. Ontario takes a whitelist approach to bait under 

which all unlisted species (including Phenacobius mirabilis) are prohibited to use as bait

(Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

There are no known physical control methods for this species.

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides (GLMRIS 2012). There are no available studies of their effects 

on suckermouth minnow at the time of this writing. 

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the US, not for use 

as euthanasia (Clearwater et al. 2008). Exposure to NaHCO3 concentration of 142-642 mg/L for 

5 min. is sufficient to anaesthetize most fish (Clearwater et al 2008). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Proterorhinus semilunaris (Heckel, 1837)

Regulations

In Canada, tubenose goby is listed as an invasive species under Canadian Federal Statutes and 

Regulations, and is thus prohibited from being possessed, released, or used as bait without a 

license. In Quebec, aquarium fish keeping, production, keeping in captivity, breeding, stocking, 

transport, sale, or purchase of live tubenose goby is prohibited by . 

In the commonwealth of Pennsylvania, it is unlawful to possess, sell, introduce, or import 

tubenose goby under . Sale, purchase, or barter of injurious, nonnative species, including 

tubenose goby, is prohibited by , and transportation in or through the commonwealth is 
196

prohibited by . In the state of Ohio, it is unlawful for a person to possess, import, or sell live 

tubenose goby under . In the state of Michigan, tubenose goby is a prohibited species under . 

Tubenose goby is regulated as an exotic fish in the state of Indiana under , meaning an individual 

must not import, possess, propagate, buy, sell, barter, trade, transfer, loan, or release into public 

or private waters any tubenose goby, including recently hatched or juvenile live fish or their 

viable eggs or genetic material. In the state of Illinois, tubenose goby is an injurious species 

under . It is unlawful to possess, propagate, buy, sell, barter, or offer to buy, sell, barter, 

transport, trade, transfer, or loan tubenose goby to any person or institution without a permit in 

Illinois. Tubenose goby is a restricted invasive species in Wisconsin under . In the state of 

Minnesota, tubenose goby is a prohibited invasive species as defined in . Ontario takes a 

whitelist approach to bait under which all unlisted species (including Proterorhinus semilunaris) 

are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

The USACE Great Lakes and Mississippi River Interbasin Study notes the potential 

effectiveness of acoustic fish deterrents in controlling or deterring Proterorhinus semilunaris 

populations (GLMRIS 2012). Acoustic deterrents include continuous wave and pulsed wave 

technology, which use sound/pressure waves to influence the behavior of aquatic organisms. 

Similarly, sensory deterrent systems such as acoustic air bubble curtains, electric barriers, and 

underwater strobe lights may prove useful in controlling populations in waterways and small 

bodies of water, but there are no studies of their effects on P. semilunaris at the present time 

(GLMRIS 2012). When using physical deterrents as barriers, combining methods can increase 

effectiveness, as was the case for Patrick et al. (1985), who found that pelagic estuarine and 

freshwater fishes were successfully deterred by a barrier combining air bubbles and strobe lights. 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides, but there are no studies of the effects of chemical treatment on 

P. semilunaris at the present time (GLMRIS 2012). 

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the US, not for use 

as euthanasia (Clearwater et al. 2008). Exposure to NaHCO3 concentration of 142-642 mg/L for 

5 min. is sufficient to anaesthetize most fish (Clearwater et al 2008). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 
197

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

Salmo trutta Linnaeus, 1758

Regulations

In Wisconsin, brown trout is restricted as a nonnative fish species in the aquaculture industry. 

Ontario takes a whitelist approach to bait under which all unlisted species (including Salmo 

trutta) are prohibited to use as bait (Ontario Fishing Guide 2014).

Control

Biological

There are no known biological control methods for this species. 

Physical

Barriers can be constructed and/or natural barriers augmented to prevent upstream migration and 

aid management and eradication efforts, though little research exists on effective barriers for 

Salmo trutta. Lintermans and Raadik (2003) noted 3 key aspects of successful barriers in relation 

to a Rainbow Trout control program: a 1.5 m or greater vertical drop; direction of water flow 

towards the middle in higher flows with no slower overland flow passing down the banks; and no 

deep pool below the barrier from which fish could jump. 

The United States Army Corps of Engineers Great Lakes and Mississippi River Interbasin Study

(USACE GLMRIS) notes the potential effectiveness of sensory deterrent systems in providing 

barriers to fish migration or eliciting fish movements, including underwater strobe lights, 

acoustic air bubble barriers, continuous and pulsed wave acoustic deterrents, and electric barriers 

(USACE 2012b). Most large-scale strobe systems consist of four individual lights that flash at a 

rate of 450 flashes/minute, with an approximate intensity of 2634 lumens/flash (USACE 2012b).

The United States National Park Service uses physical removal through electrofishing to manage 

Rainbow Trout and Brown Trout populations that threaten native Brook Trout in Shenandoah 

National Park, Virginia (NPS 2011). 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides (USACE 2012b).

In 1995, a total of 20 km of stream length (seven different streams) was treated in Victoria, 

Australia with a total of 60 L rotenone used, neutralized with 1100 kg of potassium 

permanganate, and with rotenone volumes ranging from 0.3 to 0.5 L per 100 m of stream, in 

order to eradicate Rainbow Trout (Oncorhynchus mykiss) and Brown Trout and protect native 

Galaxiids (Lintermans and Raadik 2003). Areas in which Trout and Galaxiids overlapped were 

not treated with Piscicides; they were instead intensively electrofished to more selectively 

remove Trout (Lintermans and Raadik 2003). Rotenone has also been used to eradicate Brown 

Trout in two streams of Kaiwharawhara catchment in Wellington, New Zealand using a 

concentration of 200 μg/L rotenone over initial treatment times of 4 and 5.5 hours for the smaller 
198

and larger stream, respectively (Pham et al. 2013). Ling (2003) noted that Brown Trout has an 

LC50 of 5.5 μg/L at 17°C for 1h exposure to rotenone.

Boogaard et al. (2003) found that Brown Trout are among the least sensitive fishes to the 

lampricide TFM and a TFM/1% niclosamide mixture in laboratory exposures, with a 12 hour

LC50 of 8.90 mg/L for TFM and 12 hour LC50s of 5.01 and 5.68 mg/L for TFM/1% 

niclosamide. Sensitivity was slightly decreased in field tests (Boogaard et al. 2003).

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). 

Salmonids are considered to be among the most sensitive fishes to low dissolved oxygen levels, 

with a DO concentration of 1-3 mg/L sufficient to cause mortality or loss of equilibrium 

(Clearwater et al. 2008). However, CO2 is approved only for use as an anesthetic for cold, cool, 

and warm water fishes the United States, not for use as euthanasia (Clearwater et al. 2008). 

Exposure to NaHCO3 concentration of 142-642 mg/L for five minutes is sufficient to 

anaesthetize most fish (Clearwater et al. 2008). 

Low pH is known to affect fish behavior. Ikuta et al. (2003) documented the effects of low pH on 

S. trutta, noting that trout would not swim upstream into areas of pH lower than 5.5. Acute

exposure to low pH levels can directly kill fish by discharge of sodium and chloride ions from 

body fluid, and sub-lethal levels can affect reproduction (Ikuta et al. 2003). In the case of Brown 

Trout, weak acidic conditions of <pH 6.4 were enough to depress prespawning digging behavior. 

Salmonids in general show significant avoidance of acidic environments (Ikuta et al. 2003). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Boogaard et al. (2003) found 

that the lampricides 3-trifluoromethyl-4-nitrophenol (TFM) and 2’,5-dichloro-4’-

nitrosalicylanilide (niclosamide) demonstrate additive toxicity when combined. In another study 

on cumulative toxicity, combinations of Bayer 73 (niclosamide) and TFM with contaminants 

common in the Great Lakes (pesticides, heavy metals, industrial organics, phosphorus, and 

sediments) were found to be mostly additive in toxicity to Rainbow Trout, and one combination 

of TFM, Delnav, and malathion was synergistic, with toxicity magnified 7.9 times (Marking and 

Bills 1985). This highlights the need for managers to conduct on-site toxicity testing and to give 

serious consideration to determining the total toxic burden to which organisms may be exposed 

when using chemical treatments (Marking and Bills 1985). Other non-selective alterations of 

water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 
199

Scardinius erythrophthalmus Linnaeus, 1758

Regulations (pertaining to the Great Lakes region)

In Quebec, aquarium fish-keeping, production, keeping in captivity, breeding, stocking, 

transport, sale or purchase of live Rudd is prohibited (Quebec Statutes and Regulations RRW § c 

C-61.1, r7). In Ontario, Rudd is listed as an invasive fish (Ontario Fishery Regulations § 

SOR/2007-237) and its use as bait is prohibited. In Pennsylvania, it is unlawful to possess, 

import, or introduce Rudd (58 PA Code § 71.6). In Ohio, it is unlawful to possess, import, or sell 

Rudd (Ohio Administrative Code § 1501:31-19). In Michigan, Rudd is a prohibited fish species 

(MI NREPA 451 § 324.41301). In Indiana, it is unlawful to import, possess, propagate, buy, sell, 

barter, trade, transfer, loan, or release into public or private waters any adult or recently hatched 

or juvenile Rudd or their genetic material (312 IN Administrative § 9-6-7). In Illinois, Rudd is an 

injurious species, meaning it shall not be possessed, propagated, bought, sold, bartered or offered 

to be bought, sold, bartered, transported, traded, transferred or loaned to any other person or 

institution unless a permit is first obtained from the Department of Natural Resources in 

accordance with Illinois Administrative Code § 805.40, except persons engaged in interstate 

transport for lawful commercial purposes who do not buy, sell, barter, trade, transfer, loan or 

offer to do so in Illinois (IL Administrative Code § 805). In Wisconsin, Rudd is a prohibited

species under Wisconsin Administrative Code § NR 40.04, meaning that no person may 

transport, possess, transfer, or introduce Rudd, except as otherwise provided in paragraphs (b) to 

(h) of Wisconsin Administrative Code § NR 40.04. In Minnesota, Rudd is a prohibited species, 

meaning it is unlawful (a misdemeanor) to possess, import, purchase, transport, or introduce 

except under a permit for disposal, control, research, or education (MN Administrative Rules § 

6216.0250).

Control

Biological

There are no known biological control methods for this species. 

Physical

Fine-mesh monofilament gill nets have been used to control Rudd in three shallow lakes in 

Waikato, New Zealand, but elimination was not achievable (Neilson et al. 2004). Small, 

potentially fecund fish in dense littoral vegetation proved challenging to net, presenting a 

problem for total eradication, but removal of larger Rudd likely affected breeding success and 

netting is seen as a highly cost effective control method with low environmental impact (Neilson 

et al. 2004). 

Chemical

Of the four chemical piscicides registered for use in the United States, antimycin A and rotenone 

are considered general piscicides (USACE 2012b). Ling (2003) noted that rotenone has an LC50 

of 24.5 μg/L for one hour exposure to Rudd at 20°C. 

Increasing CO2 concentrations, either by bubbling pressurized gas directly into water or by the 

addition of sodium bicarbonate (NaHCO3) has been used to sedate fish with minimal residual 

toxicity, and is a potential method of harvesting fish for removal, though maintaining adequate 

CO2 concentrations may be difficult in large/natural water bodies (Clearwater et al. 2008). CO2 is 

approved only for use as an anesthetic for cold, cool, and warm water fishes the United States, 
200

not for use as euthanasia (Clearwater et al. 2008). Exposure to NaHCO3 concentration of 142-

642 mg/L for five minutes is sufficient to anaesthetize most fish (Clearwater et al. 2008). 

It should be noted that chemical treatment will often lead to non-target kills, and so all options 

for management of a species should be adequately studied before a decision is made to use 

piscicides or other chemicals. Potential effects on non-target plants and organisms, including 

macroinvertebrates and other fishes, should always be deliberately evaluated and analyzed. The 

effects of combinations of management chemicals and other toxicants, whether intentional or 

unintentional, should be understood prior to chemical treatment. Other non-selective alterations 

of water quality, such as reducing dissolved oxygen levels or altering pH, could also have a 

deleterious impact on native fishes, invertebrates, and other fauna or flora, and their potential 

harmful effects should therefore be evaluated thoroughly. 

A.11 Insects

Acentria ephemerella Olivier, 1791

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

Acentria ephemerella is used for biological control of Eurasian Watermilfoil (Myriophyllum 

spicatum). Its population is best controlled by elimination of its host plants, which are 

predominantly Eurasian Watermilfoil but may to a lesser extent, include a variety of other native 

and nonindigenous plants (Cornell 2004).

Physical

Mechanical harvesting, herbicide applications, benthic barriers, and water drawdowns all remove 

either A. ephemerella individuals or their habitat (aquatic plants) from waterways (Cornell 

2004).

Chemical

This species is susceptible to herbicide control of its host plants (Cornell 2004).

Tanysphyrus lemnae Paykull/Fabricius, 1792

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological, physical or chemical control methods for this species.
201

A.12 Mollusks

Corbicula fluminea O. F. Müller, 1774

Regulations (pertaining to the Great Lakes region)

Corbicula fluminea is a prohibited species in Wisconsin (Wisconsin Administrative Code § NR 

40.04). In Indiana an individual may not import, possess or release Asiatic clams (312 IN 

Administrative Code § 9-9-3 and § 14-22-17-3). 

Control

Eradication of Asian Clams from infested open waters is unlikely – emphasis is generally on 

preventing further spread.

Biological 

There are no known biological control methods for this species.

Physical

Screens and traps are commonly employed to prevent C. fluminea colonization of water intakes 

(GISD 2013). 

Diver assisted suction removal and bottom barriers are being researched as potential methods for 

physical control of C. fluminea populations in Lake Tahoe (UC Davis TERC, 2004). Benthic 

barriers have been demonstrated to be effective for short-term control of C. fluminea, but nontarget mortality to other benthic invertebrates may be high (Wittmann et al. 2012).

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N,N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

Low concentration of chlorine or bromine will kill juvenile Asian Clams (GISD 2013). 

C. fluminea is not tolerant of fluctuating environmental conditions (particularly temperature and 

oxygen) and is prone to massive die-offs (Menninger 2013), this suggests that short-term 

chemical manipulation may be useful in controlling C. fluminea populations. C. fluminea may be 

controlled at intake pipes by heating influent water to 37oC (GISD 2013). 
202

Dreissena polymorpha Pallas, 1771

Regulations (pertaining to the Great Lakes region)

Federal law (Lacey Act 1990) prohibits the possession and transportation of Zebra Mussels in the 

United States unless intended for research. 

The following regulations apply to all vessels equipped with ballast water tanks that enter a 

United States port on the Great Lakes after operating in waters beyond the exclusive economic 

zone. Vessels are required to exchange ballast water beyond the exclusive zone prior to entering 

any Great Lakes port. Ballast exchange may also be conducted in waters that are considered a 

non-threat to the infestation of an aquatic nuisance species in the Great Lakes (Allowable waters 

stated under ANS Task Force section 1102(a)(1)). Vessels are also required to use 

environmentally sound ballast water management methods if deemed necessary (NASPCA 

2000).

In New York, it is unlawful to intentionally release Zebra Mussels into state waters (New York 

ECL § 11-0507). In Pennsylvania, it is unlawful to possess, introduce, import, transport, sell, 

purchase, offer for sale, or barter Zebra Mussels (58 PA Code § 63.46, 71.6, and 73.1). In Ohio, 

it is unlawful to possess, import, or sell Zebra Mussels (Ohio Administrative Code § 1501:31-19-

01(K)(4)). In Indiana, it is unlawful to import, possess, or release Zebra Mussels into public or 

private waters (312 IN Administrative Code § 9-9-3(d) and (e)). In Michigan, Zebra Mussels are 

a restricted species (MI Compiled Laws § 324.41301) and therefore cannot be possessed unless it 

is to identify, eradicate, or control the species (MI Compiled Laws § 324.41303). In Wisconsin, 

it is unlawful to transport, transfer, or introduce Zebra Mussels (WI Administrative Code § NR 

40.05). In Minnesota, it is unlawful to place or attempt to place a watercraft, trailer, or plant 

harvesting equipment that has Zebra Mussels attached into state water (MN Statutes § 84D.10). 

Persons leaving the state are required to drain boats and related equipment during transportation 

on a public road (MN Statutes § 84D.10, MN Administrative Rules 6216.0500, Kaminski Leduc 

2011). Illinois lists Zebra Mussels as injurious species (IL Administrative Code § 805).

Control

Given the widespread established of Zebra Mussels in the Great Lakes, total eradication is 

considered impossible with current technologies. No control methods are currently available for 

open water applications. Control efforts focus primarily on protection of human infrastructure 

(such as water intakes) and along vectors of spread (such as boats, trailers, gear, etc). Controlling 

Zebra Mussels to minimize effect on natural and anthropologic systems is expensive, regardless 

of the method(s) chosen. 

Biological 

Biological control so far has proven to be ineffective in controlling Dreissena species. Predation 

by migrating diving ducks, fish species, and crayfish may reduce mussel abundance, though the 

effects are short-lived (Bially and MacIsaac 2000). Other biological controls being researched 

are selectively toxic microbes and parasites that may play a role in management of Dreissena

populations (Molloy 1998). Laboratory testing shows strain CL145A of Pseudomonas 

fluorescens (a bacterium) to be highly lethal to Zebra Mussels; capable of eliminating over 90% 

of adults and 100% of larvae (Molloy 2002, Abdel-Fattah 2011). Commercially, this product is 

known as Zequanox® and is developed by Marrone Bio Innovations (Abdel-Fattah 2011).

Interfering with the synchronization of spawning by adults in their release of gametes could also 
203

offer control of Dreissena populations (Snyder et al. 1997). Another approach would be to 

inhibit the planktonic veliger (larvae) from settling and attaching to a surface to begin 

development (Kennedy 2002). 

Physical

Effective physical controls of Dreissena include drawing water from public sources or a 

groundwater well, infiltration intakes or sandfilter intakes (filter out veligers), thermal 

treatments, carbon dioxide pellet blasting, high-pressure water jet cleaning, mechanical cleaning, 

freezing, scraping, scrubbing, pigging, and desiccation. Potential controls include the use of 

electrical fields, pulse acoustics, low-frequency electromagnetism, ultraviolet light (UV light), 

and reduced pressure (USACE 2002).

Physical removal of visible vegetation (which may harbor small mussels) from boats, trailers and 

other equipment being moved from one water body to another is an important method in 

controlling the spread of Zebra Mussels. Flushing engines, cooling systems, live wells and bilge 

with water over 43oC will kill veligers and 60oC will kill adults. Air drying equipment for five

days will kill most larvae and smaller mussels, but large mussels may survive two weeks out of 

water.

Placement of water intakes (in areas too deep or otherwise unsuitable for Zebra Mussel

colonization) has been used as a form of physical control, but this is less successful in areas 

which also have Quagga Mussels.

Chemical

Oxidizing chemical control treatments effective against D. polymorpha include hypochlorite 

reaction, chloramine, chlorine dioxide, bromine, ozone, potassium, permanganate, and sodium 

chlorite. Non-oxidizing chemicals include molluscicides, detoxification, and potassium ions. 

However, application of these chemical can be extremely detrimental to ecosystem and human 

health so possible effects should be thoroughly evaluated before use (Boelman et al. 1997, 

Sprecher and Getsinger 2000).

Various chemical coatings – including copper-based, tributyltin-based, copolymer, vinyl/epoxy, 

resin and other films - can be applied to structures to prevent the attachment of Zebra Mussels. 

Tributyltin-based antifoulants are extremely toxic and restricted by federal law (Ohio Sea Grant 

1992). 

Dreissena rostriformis bugensis Andrusov, 1897

Regulations (pertaining to the Great Lakes region)

Federal law (Lacey Act 1990) prohibits the possession and transportation of Quagga Mussels in 

the United States unless intended for research. 

The following regulations apply to all vessels equipped with ballast water tanks that enter a 

United States port on the Great Lakes after operating in waters beyond the exclusive economic 

zone. Vessels are required to exchange ballast water beyond the exclusive zone prior to entering 

any Great Lakes port. Ballast exchange may also be conducted in waters that are considered a 

non- threat to the infestation of an aquatic nuisance species in the Great Lakes (Allowable waters 

stated under ANS Task Force section 1102(a)(1)). Vessels are also required to use 
204

environmentally sound ballast water management methods if deemed necessary (NASPCA 

2000). 

In Pennsylvania, it is unlawful to possess, introduce, import, transport, sell, purchase, offer for 

sale, or barter Quagga Mussels (58 PA Code § 63.46, 71.6, and 73.1). In Indiana, it is unlawful to 

import, possess, or release Quagga Mussels into public or private waters (312 IN Administrative 

Code § 9-9-3(d) and (e)). In Michigan, Quagga Mussels are a restricted species (MI Compiled 

Laws § 324.41301) and therefore cannot be possessed unless it is to identify, eradicate, or control 

the species (MI Compiled Laws § 324.41303). In Wisconsin, it is unlawful to transport, transfer, 

or introduce Quagga Mussels (WI Administrative Code § NR 40.05). In Minnesota, it is 

unlawful to place or attempt to place a watercraft, trailer, or plant harvesting equipment that has 

Quagga Mussels attached into state water (MN Statutes § 84D.10). Persons leaving the state are 

required to drain boats and related equipment during transportation on a public road (MN 

Statutes § 84D.10, MN Administrative Rules 6216.0500, Kaminski Leduc 2011). Violation 

penalties can range from a civil fine of $250-$1,000 and/or a misdemeanor (MN Statutes § 

84D.13) Illinois lists Quagga Mussels as injurious species (IL Administrative Code § 805).

Control

Given the widespread established of quagga mussels in the Great Lakes, total eradication is 

considered impossible with current technologies. No control methods are currently available for 

open water applications. Control efforts focus primarily on protection of human infrastructure 

(such as water intakes) and along vectors of spread (such as boats, trailers, gear, etc). Controlling 

quagga mussels to minimize effect on natural and anthropologic systems is expensive, regardless 

of the method(s) chosen.

Biological 

Biological control so far has proven to be ineffective in controlling Dreissena species. Predation 

by migrating diving ducks, fish species, and crayfish may reduce mussel abundance, though the 

effects are short-lived (Bially and MacIsaac 2000). Other biological controls being researched 

are selectively toxic microbes and parasites that may play a role in management of Dreissena

populations (Molloy 1998). Laboratory testing shows strain CL145A of Pseudomonas 

fluorescens (a bacterium) to be highly lethal to quagga mussels; capable of eliminating over 90% 

of adults and 100% of larvae (Abdel-Fattah 2011, Molloy 2002). Commercially, this product is 

known as Zequanox® and is developed by Marrone Bio Innovations (Abdel-Fattah 2011).

Interfering with the synchronization of spawning by adults in their release of gametes could also 

offer control of Dreissena populations (Snyder et al. 1997). Another approach would be to 

inhibit the planktonic veliger (larvae) from settling and attaching to a surface to begin 

development (Kennedy 2002). 

Physical

Effective physical controls of Dreissena include drawing water from public sources or a 

groundwater well, infiltration intakes or sandfilter intakes (filter out veligers), thermal 

treatments, carbon dioxide pellet blasting, high-pressure water jet cleaning, mechanical cleaning, 

freezing, scraping, scrubbing, pigging, and desiccation. Potential controls include the use of 

electrical fields, pulse acoustics, low-frequency electromagnetism, ultraviolet light (UV light), 

and reduced pressure USACE 2002).
205

A recent study has confirmed that thermal treatment of residual water in boats and other water 

vehicles may be a viable option for managing quagga mussel spread. Increasing temperature and 

exposure time was found to increase the level of veliger mortality (Craft and Myrick 2011). 

Other methods of physical control include exposure and desiccation, manual scraping, highpressure jetting (including with high temperature water), mechanical filtration, removable 

substrates, and sonic vibration. 

Chemical

Prechlorination has been the most common treatment for control of Dreissena mussels, but 

chlorine concentrations needed for effective control of quagga mussels may reach hazardous 

levels (Grime 1995). Potassium permanganate has been used as an alternative control, especially 

for drinking water sources. Ozone is also a potential control. Other molluscides and anti-fouling 

coatings may be effective; however, application of these chemical can be extremely detrimental 

to ecosystem and human health so possible effects should be thoroughly evaluated before use 

(Boelman et al. 1997, Sprecher and Getsinger 2000).

Other

Other potential methods of Quagga Mussel control include oxygen deprivation, radiation, and 

electric currents.

Lasmigona subviridis Conrad, 1835

Regulations (pertaining to the Great Lakes region)

NatureServe lists this species as ‘global vulnerable’ at moderate risk of extinction in its native 

range. 

There are no regulations for this species as an invasive in the Great Lakes region.

Control

Management research focuses primarily on preservation of populations within the native range. 

Biological 

Competition with dreissenid mussels will likely limit its expansion in the Great Lakes.

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).
206

Pisidium amnicum Müller, 1774

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

Pisidium henslowanum Shepard, 1825

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 
207

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

Pisidium moitessierianum Paladilhe, 1866

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Research on drawdown as a potential control technology for P. moitessierianum suggests that it 

will be in-effective as a control in most situations (Mouthon 2011). Pisidiid clams showed poor 

resistance to a brief period of drying of the habitat, but also demonstrated high rates of recovery. 

In the study sites, drawdown did cause a decline in P. moitessierianum, but the population 

recovered quickly, becoming the dominant bivalve in the following year. 

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

Pisidium supinum Schmidt 1850

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.
208

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

Sphaerium corneum Linnaeus, 1758

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

A wide array of chemical molluscicides are available, but are not species-specific and may harm 

native species to a greater extent than non-natives. 

Molluscicides are typically classified as either oxidizing or non-oxidizing compounds. Oxidizing 

chemicals include chlorine, chlorine dioxide, chloramines, ozone, bromine, hydrogen peroxide, 

and potassium permanganate. Non-oxidizing chemicals (including organic film-forming 

antifouling compounds, gill membrane toxins, and nonorganics) can be classified into several 

distinct groups: quanternary and polyquaternary ammonium compounds; aromatic hydrocarbons; 

endothall as the mono (N, N-dimethylalkyl amine) salt; metals and their salts (e.g., copper sulfate 

formulations); and niclosamide (including some formulations of Bayluscide). Bayluscide was 

initially developed as a Sea Lamprey larvicide, but has molluscicidal activity. While some of 

these products are biodegradable, many require detoxification or deactivation to meet state and 

Federal discharge requirements (USACE 2012a).

In short-term experiments, S. corneum can reduce the bioaccumulation of 2, 4, 5-trichlorophenol 

(TCP) by closing their shell valves (Heinonen et al. 1997) – this reaction to chemical stimuli 

generally may limit the usefulness of chemical molluscicides against this species.
209

Bithynia tentaculata

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are currently no documented successful methods for the control of Faucet Snails in open 

water ecosystems. 

Biological 

There are no known biological control methods for this species.

Physical

In an attempt to limit the number of Faucet Snails in the Upper Mississippi River National 

Wildlife and Fish Refuge, biologists experimented with covering colonies of Faucet Snails with 

sand (Williams 2007). The success of this method was undocumented.

Chemical

There are no known chemical control methods for this species.

Cipangopaludina chinensis malleata

Regulations (pertaining to the Great Lakes region)

Chinese Mysterysnail is a regulated invasive species in Minnesota (MN Administrative Rules § 

6216.0260) and a restricted species in Wisconsin (Wisconsin Administrative Rules § 40.05). 

Control

Specific control methods for the Chinese Mysterysnail have yet to be developed.

Biological 

Manipulation of predator fishes and turtles that eat snails may be useful in the control of snail 

populations. However, as a relatively large snail species, Cipangopaludina chinensis may escape 

predation by smaller fishes. 

Physical

Preliminary research demonstrates that C. chinenesis will not migrate upstream against a small 

current (Rivera 2008). Authors suggest that acceleration of current may be an important 

management tool for preventing upstream spread.

Dessication (drying) is not an effective control method for C. chinensis. Field experiments under 

mesic conditions indicated that this snail can survive exposure to air for at least four weeks 

(Havel 2011).

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill. With Chinese Mysterysnails possessing the ability to “close up”, more damage 

would probably occur to native snails in the treatment area than to the target pest. 
210

Cipangopaludina japonica

Regulations (pertaining to the Great Lakes region)

Japanese Mysterysnail is a regulated invasive species in Minnesota (MN Administrative Rules §

6216.0260)

Control

Specific control methods for the Japanese Mysterysnail have yet to be developed – most 

available control suggestions are based on research with the closely related Cipangopaludina 

chinensis.

Biological 

Manipulation of predator fishes and turtles that eat snails may be useful in the control of snail 

populations. However, as a relatively large snail species, Cipangopaludina japonica may escape 

predation by smaller fishes. 

Physical

Preliminary research demonstrates that C. chinenesis will not migrate upstream against a small 

current (Rivera 2008) – it is not known whether C. japonica is similarly restricted. Acceleration 

of current may be an important management tool for preventing upstream spread.

Dessication (drying) is not likely to be an effective control method for C. japonica. Field 

experiments under mesic conditions indicated that C. chinensis and other snails with opercula 

can survive exposure to air for at least four weeks (Havel 2011).

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill. With Japanese Mysterysnails possessing opercula, more damage would probably 

occur to native snails in the treatment area than to the target pest. 

Elimia virginica

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Specific control methods for Elimia virginica have yet to be developed.

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill. With E. virginica possessing the ability to close their shells (opercula), more 

damage would probably occur to native snails in the treatment area than to the target pest. 
211

Gillia altilis

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Specific control methods for Gillia altilis have yet to be developed.

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill.

Potamopyrgus antipodarum

Regulations (pertaining to the Great Lakes region)

New Zealand Mudsnails are listed as a prohibited species in Wisconsin (Wisconsin 

Administrative Rules § 40.05) and Minnesota (MN Administrative Rules § 6216.0260).

Control

Many times New Zealand Mudsnails may be in a river or lake where chemical eradication will 

not be feasible and physical eradication difficult. Areas where eradication may be possible 

include small lakes and ponds, waterbodies that can be temporarily hydrologically separated. 

Biological 

Parasites of New Zealand Mudsnails fromNew Zealand may also become useful to control 

population size by inhibiting reproduction. Studies of the efficacy and specificity of a trematode 

parasite from the native range of New Zealand Mudsnails as a biological control agent have 

shown positive results so far (Dybdahl et al. 2005).

Physical

New Zealand Mudsnails easily hitchhike with fish and aquatic plants. Inspection of 

boats/trailers/gear is essential, but equipment should also be dried thoroughly before moving 

from infected to uninfected waters. Putting fishing gear in a freezer for 6-8 hours will kill all 

attached New Zealand Mudsnails (Medhurst 2003, Richards et al. 2004). Putting fishing gear in 

water maintained at 49°C for a few minutes will eliminate New Zealand Mudsnails (Medhurst 

2003). Mudsnails can survive at 43.3°C so the water temperature needs to be accurate. Dry 

fishing gear at 28-30°C for at least 24 hours or at 40°C for at least two hours (Richards et al. 

2004). 

For (aquaculture) facilities where no known New Zealand Mudsnail contamination occurs, close 

visual inspection of water systems, raceways, stocking equipment, as well as regular gut content 

analysis can detect the arrival of snails before they can be spread . 
212

Physical treatments include the use of temperature, humidity or desiccation to kill the target 

species. This includes draining the infested areas. New Zealand Mudsnails can survive for long 

periods in a cool damp environment; however, draining the areas where they are congregated and 

exposing them to sunlight during the summer months may be sufficient for eradication. Using a 

flame thrower in a hatchery situation against the walls of raceways will kill any mudsnails 

attached. Mudsnails cannot withstand warm temperatures (Dwyer et al. 2003, Richards et al. 

2004) or low humidity situations (Dwyer and Kerans, unpublished, Richards et al. 2004). 

Alternately, if an infested area could be drained in the winter and the substrate is frozen to a 

depth containing the mudsnails, then total eradication will occur. There is preliminary evidence 

that hydrocyclonic separators may also be a useful tool to decontaminate fish hatchery water 

supplies and prevent the spread of New Zealand Mudsnails within a hatchery. 

It has been suggested that barriers such as copper stripping or electrical weirs may limit 

volitional movement of New Zealand Mudsnails, particularly as a means of protecting high risk 

sites like fish hatchery water systems. Some investigations are underway but there is no 

applicable tool available yet. 

Chemical

Chemical methods used to eradicate New Zealand Mudsnails include: Bayer 73, copper sulfate, 

and 4-nitro-3-trifluoromethylphenol sodium salt (TFM). The only molluscicide known to have 

been tested against New Zealand Mudsnails is Bayluscide (niclosamide). Preliminary 

investigations also suggest that copper and carbon dioxide under pressure may prove useful in 

both decontaminating fish hatchery water supplies and preventing spread into uncontaminated 

areas of a hatchery. Ozone has not been shown to be effective in killing New Zealand Mudsnails 

in a hatchery environment. 

The most effective solutions for killing New Zealand Mudsnails which can be used in the field, 

according to this research are copper sulfate (252mg/L Cu), benzethonium chloride (1,940 mg/L) 

and 50% Commercial Solutions Formula 409® Cleaner Degreaser Disinfectant. 

Copper sulfate, hyamine and hydrogen peroxide, have all been used to control New Zealand mud 

snails (IJC 2011). 

Radix auricularia

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Specific control methods for Radix auricularia have yet to be developed.

Biological 

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.
213

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill.

Valvata piscinalis

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Specific control methods for Valvata piscinalis have yet to be developed.

Biological 

Manipulation of predator fishes and turtles that eat snails may be useful in the control of snail 

populations. However, as a relatively large snail species, V. piscinalis may escape predation by 

smaller fishes. 

Physical

There are no known physical control methods for this species.

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill. With V. piscinalis possessing the ability to “close up”, more damage would 

probably occur to native snails in the treatment area than to the target pest. 

Viviparus georgianus I. Lea, 1834

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Specific control methods for Viviparus georgianus have yet to be developed.

Biological 

Manipulation of predator fishes and turtles that eat snails may be useful in the control of snail 

populations. However, as a relatively large snail species, V. georgianus may escape predation by 

smaller fishes. 

Physical

There are no known physical control methods for this species.

Chemical

There are copper compounds that are sold as snailicides but they are usually not selective in the 

snails they kill. With V. georgianus possessing the ability to “close up”, more damage would 

probably occur to native snails in the treatment area than to the target pest. 
214

A.13 Plants

Agrostis gigantea Roth

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. However, the Michigan Department of Natural 

Resources (MIDNR and MIDEQ 2009) recommends use of Redtop for erosion control in 

forested land. Moreover, Redtop’s close cousin, Creeping Bentgrass (A. stolonifera), is 

commonly put to commercial use on golf course putting greens, tees, and fairway turf (Banks et 

al. 2004).

Control

Biological

There are no known biological control methods for this species.

Physical

This species has rhizomal roots that can grow to a depth of four feet (Tilley et al. 2010), making 

mechanical removal of Redtop difficult, time-consuming, and unlikely to be successful, as any 

significant portions of the rhizome that are not removed from the soil could generate new plants.

Continuous grazing or mowing to a height of less than three inches may result in temporary 

control (Tilley et al. 2010).

Chemical

Due to the tendency for this species to form dense, monospecific turfs (Tilley et al. 2010) 

herbicide application should be easy to manage. Increased grass chlorosis (yellowing of the plant 

due to reduced chlorophyll production) results when herbicides are applied during cool weather 

(10°C) (McCullough and Hart 2006).

Among available herbicides, Redtop is very susceptible to atrazine (Carey 1995). It can also be 

well controlled by glyphosate. However, glyphosate-resistant strains of the related Creeping 

Bentgrass (A. stolonifera) do exist and are becoming more common (Hart et al. 2005); 

commercial release is likely to increase the potential for unintended transfer of the herbicide 

resistance gene to A. gigantea (NatureServe 2008).

Given the ease of hybridization with other Agrostis species (Tilley et al. 2010), if control 

methods listed above are not successful, alternative chemical control methods may also be 

considered. For instance, in tests conducted in Virginia, the closely related Creeping Bentgrass

experienced at least a 92% die-off after applying isoxaflutole, imazaquin, or mesotrione in two to 

three sequential applications (Beam et al. 2006). Fluazifop-P has also been effective in 

controlling bentgrass species (Hart et al. 2005). In addition, Creeping Bentgrass seems to be 

susceptible to rimsulfuron, especially when the herbicide is kept dry after application (Barker et 

al. 2005).
215

Alnus glutinosa (L.) Gaertn. 

Regulations (pertaining to the Great Lakes region)

There are conflicting recommendations on the regulation of Black Alder. Indiana recommends 

that this species be inter-planted to improve soil quality and “protect other valuable trees” (IDNR 

n.d.). The state also includes Black Alder on a list of invasive exotics plants whose “use in 

landscaping and re-vegetation projects should be avoided or limited when possible” (Homoya 

2010). Black Alder is also listed as a “plant to avoid” in Wisconsin’s planting guide (WIDNR 

n.d.). It is a recommended tree for urban environments in Minnesota (Johnson and Himanga 

2009).

Control

For the most effective control, the United States Department of Agriculture Natural Resources 

Conservation Service (2006) recommends treating Black Alder with a combination of physical 

and chemical methods.

Biological

There are no known biological control methods for this species.

Physical

Trees should be felled and then herbicide applied to the stumps to prevent sprouting (USDA 

NRCS 2006).

Chemical

Effective herbicides include napropamide (preemergence) (Willoughby et al. 2007), glyphosate 

via exposed stump application (Kelly and Southwood 2006) triclopyr triethylamine via foliar 

application (Champion et al. 2008).

Alopecurus geniculatus L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

Toogood et al. (2008) found that Water Foxtail tolerates periods of waterlogging and flooding. 

This species does not grow as well under a drier water regime. Moreover, in locations where 

water level manipulation is used for wetland management, flooding should not be used as a 

method for controlling this species.

Alopecurus geniculatus has the ability to reproduce via seed dispersal and vegetatively from 

roots nodes (Klinkenberg 2010). Each individual seed has a mass of approximately 0.8 mg, 

which is light enough to be transported short distances by wind (Klein 2011). If seeds are driven 
216

below this depth, they are unlikely to germinate. A variety of tilling methods can be used both to 

kill seedlings after emergence and to bury seeds deep into the soil to prevent germination 

(Curran 2009). Seeds of Water Foxtail can remain viable in soil for at least three years (Roberts 

1986). Therefore, management activity would be most effective if performed before seeds have a 

chance to disperse.

Chemical

Complete eradication of a related species, Alopecurus pratensis L. (Meadow Foxtail), was 

achieved when treated in late summer with glyphosate at 1.0 kg/ha (0.89 lbs/ac). Use of fluazifop 

also controls A. pratensis L., but has also been reported to be toxic to fishes and aquatic 

invertebrates. Absorption of sethoxidon, another herbicide used to control A. pratensis L., can be 

increased when it is simultaneously applied with oil adjuvant and a non-ionic surfactant (OSU 

2005). If managers expect dense populations of foxtail species, grass herbicide rates may need to 

be increased or the application timing altered to include split applications or postemergence 

control (Curran 2009).

Butomus umbellatus L.

Regulations (pertaining to the Great Lakes region)

Butomus umbellatus is listed as a prohibited species in Michigan, Minnesota, and Illinois, and as 

a restricted species (but still available) in Wisconsin (GLPANS 2008, Jensen 2011). A recent 

survey of Minnesota Nursery and Landscape Association Members revealed that 80% of 

respondents were incorrect or unsure when judging the non-native status of B. umbellatus despite 

its prohibited use in Minnesota (Peters et al. 2006). It is on New York State’s Interim Invasive 

Species Plant List (NYSDEC 2011). In Pennsylvania it is labeled as a medium-high threat to 

native ecosystem and in Ohio it is classified as a “well established invasive plant” (Ohio 

Division of Natural Areas and Preserves and Nature Conservancy 2000, Higman and Campbell 

2009).

Control

Butomus umbellatus has a similar appearance to some native plants, such as common bulrush 

(Typha latifolia) (Jensen 2011). Care should be taken to first identify the plants in question 

before control actions are taken.

Biological

In its native range in Europe, ducks have been known to graze extensively on Flowering Rush

(Hroudová et al. 1996).

Physical

B. umbellatus spreads by floating seeds and vegetatively by rhizomes and root pieces (Campbell 

et al. 2010, Jensen 2011). Great care to remove all parts of the plant should be taken when 

implementing a physical method of control. To ensure removed plants will not sent out shoots, 

thoroughly dry all plant pieces (Jensen 2011).

Cutting Flowering Rush below the water surface will not kill the plant, but it will reduce its 

abundance, and therefore, its ability to spread. Multiple cuts may be needed throughout the 
217

growing season (Jensen 2011). Cutting later may remove most of that season’s rhizome growth 

(Hroudová et al. 1996). Hand digging may be a viable option for removing isolated plants 

(Jensen 2011). Techniques involving raking or pulling are not recommended (Jensen 2011).

Decreases in water level promotes establishment of Flowering Rush, suggesting that water level 

reductions is not a method of control (Parkinson et al. 2010).

Chemical

Control with the use of herbicides is difficult because these chemicals easily miss or fall off of 

the narrow, slightly twisted leaves of Flowering Rush (Seizer 2009, Parkinson et al. 2010). There 

is currently no herbicide that targets B. umbellatus, but initial testing shows that a mid-summer 

application of imazapyr (labeled as the herbicide ‘Habitat’) during calm weather may be 

effective (Parkinson et al. 2010, MN DNR 2012b). Some applicators report control of Butomus 

umbellatus with glyphosate; others report control of submerged B. umbellatus with diquat in sites 

that have limited dilution potential.

Cabomba caroliniana

Regulations (pertaining to the Great Lakes)

Cabomba caroliniana is prohibited in Wisconsin, Illinois and Michigan (GLPANS 2008). In 

Minnesota, C. caroliniana can be possessed, sold, bought, and transported, but it is illegal to 

release it into the environment (MN DNR 2013b). The New York Invasive Species Council 

ranks this species as posing a high ecological threat (NYISC 2010). 

The Great Lakes Indian Fish & Wildlife Commission have not found C. caroliniana in their 

ceded territories, but recommended immediate control upon detection (Falck and Garske 2003).

Control

Biological

When fed on my crayfish and snails, C. caroliniana induces a chemical defense mechanism deter 

both herbivores and microbes that typically attack plants via openings left by herbivores 

(Morrison and Hay 2011). 

Grass Carp, Ctenopharyngodon idella, will eat C. caroliniana, but it is not their preferred food 

source. 

Diploid (fertile) Grass Carp are illegal for use in some states, such as Minnesota (MN DNR 

2013a). The use of certified triploid (sterile) Grass Carp is allowed is New York and 

Pennsylvania, with the correct permits (NY DEC 2013, Shiels and Hartle n.d.)

Physical

Cabomba caroliniana becomes brittle late in the growing season. Physical control efforts should 

not be tried during this time because broken pieces can develop into new plants (IISCTC 2007).

Physical cutting and removal of C. caroliniana is most effective on large infestations in closed 

water bodies (ADEH 2003). In areas of sufficient size and depth, this can be done with floating 
218

mechanical (CSIRO Entomology 2011). However, given the low probability of removing every 

plant fragment, this method is likely to only provide nuisance relief for a few weeks (ADEH

2003). Efficacy can be improved by using tools such as a venturi dredge, which acts like a 

vacuum cleaner to C. caroliniana fanwort fragments and the root ball (WIDNR 2012d). 

Water-level drawdowns have reduced growth of C. caroliniana populations in some areas in 

southern Wisconsin (WIDNR 2012d). Extreme drying, in which the root ball dries completely, is 

needed or the plant will return (Forest Health Staff 2006b, IISCTC 2007). 

Ensuring wash-downs of boats, trailers, and other equipment can reduce the spread of Carolina 

Fanwort (IISCTC 2007). 

Chemical 

Chemical defense also indicates that the use of biocontrol agents on invasive populations of C. 

caroliniana may not be a viable approach (Morrison and Hay 2011).

Precise application of appropriate herbicides to submerged C. caroliniana can be problematic 

and should be done with great care to avoid desired species (ADEH 2003). 

Herbicides containing endothall or fluridone have been effective in controlling C. caroliniana

(ADEH 2003, Forest Health Staff 2006b). In laboratory tests, the application of diquat and 

flumioxazin (separately) resulted in a greater than 50% reduction in photosynthesis of C. 

caroliniana, however, these trials did not include field testing (Bultemeier et al. 2009).

Michigan reports Cabomba has been controlled in one lake using whole lake 20 ppb fluridone. 

Carolina Fanwort has also been reported to be sensitive to 2,4-D (Wilson et al. 1997). Michigan 

further reports Cabomba has been controlle using spot treatment of 200 ppb flumioxazin in many 

waterbodies. However, they report that Cabomba populations have not been controlled in field 

trails with diquat or endothall. 
219

Carex acutiformis Ehrh.

Regulations (pertaining to the Great Lakes region)

Based on its competitive dominance in Stony Swamp, Ottawa, Ontario, Carex acutiformis was 

identified as a high priority invasive plant by the Canada Botanical Association in 2004, ranking 

14th overall among invasive plants (Catling 2005).

Control

Biological

There are no known biological control methods for this species.

Physical

Harvesting, either mechanically or by hand, may be a viable option, depending on site 

conditions. Extreme care should be taken to remove all parts of the roots system and rhizomes to 

prevent further spread of the species. This method is unlikely to eradicate Swamp Sedge, but it 

will help to prevent population expansion. Physical control methods are most effective when 

completed before seed production (USACE 2012b).

Seeds do not germinate when they are seven cm below the soil surface (Schütz 1998). Thus, 

tilling the soil to push the existing seeds deeper into the substrate may be an effective method for 

reducing the prevalence of Swamp Sedge (Curran et al. 2009).

Carex acutiformis does not extend into open water deeper than 55 cm, and it able to cope with 

low water levels. Stands of C. acutiformis did not decrease immediately after moderate reduction 

in water level, suggesting that water level alteration may not be an effective form of control 

(Lawniczak et al. 2010).
220

Chemical

Currently, no peer-reviewed literature examines the efficacy of herbicides against Swamp Sedge. 

According to the Center for Ecology and Hydrology all sedges are susceptible to glyphosate

(CEH 2004a); an application in mid to late summer will maximize translocation and control of 

rhizomes (USACE 2012b). Imazapyr has also been effective in controlling some sedge species 

(USACE 2012b).

Carex disticha Huds.

Regulations (pertaining to the Great Lakes region)

Tworank Sedge has not been found in the United States, but it is classified as an “introduced 

species” in Ontario, Canada (Canadensys 2012, USDA NRCS 2012).

Control

Biological

There are no known biological control methods for this species.

Physical

Carex spp. are able to withstand water-logged conditions, suggesting that water level increase 

may not be an effective form of control (Riutta et al. 2007).

Chemical

There are no known chemical control methods for Carex disticha; however, imazapyr has been 

effective in controlling other sedge species (USACE 2012b).

Chenopodium glaucum

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

The occurrence of Chenopodium glaucum can be minimized in crop fields by timely spring 

tillage and early autumn plowing. Crop rotation with winter cereal grains may also be effective 

in controlling C. glaucum (Larina 2008).

Chemical

There are no known chemical control methods for this species.

Cirsium palustre

Regulations (pertaining to the Great Lakes region)

Cirsium palustre is considered to be a medium to high threat species in New York and Michigan
221

(Higman and Campbell 2009, NYISC 2010). C. palustre became a prohibited species in some 

Wisconsin counties with the creation of Wisconsin's Invasive Species Identification, 

Classification and Control Rule (Wisconsin Administrative Rules § 40.05; Terrestrial 

Herbaceous Plants Species Assessment Group 2007). Even though it is not present in Minnesota, 

it is characterized as a severe threat to native ecosystems based on its impact in other locations 

(Minnesota Invasive Species Advisory Council 2009). Marsh Thistle is listed as an introduced 

species in Ontario and Quebec (Canadensys 2012). It is considered noxious in some regions of 

British Columbia under the Weed Control Act (OLA and MAFF 2002). Marsh Thistle was 

categorized as a priority species for removal by the Great Lakes Indian Fish & Wildlife 

Commission (GLIFWC) in 2006, but this priority was reduced in subsequent comprehensive 

reports (Falck et al. 2006, 2009ab).

Control

Biological

While there are no specific biocontrol agents for C. palustre (GLIFWC 2006), herbivory by a 

variety of species may be beneficial but requires additional research.

Promising biocontrol candidates include a European seedhead fly, Terellia ruficauda (Fraser 

2000, OLA and MAFF 2002); the seed-eating weevil, Rhinocyllus conicus, currently undergoing 

experimental trial in the Robson Valley Forest District, British Columbia (OLA and MAFF 

2002, USDA Forest Service 2005b); and the glassy cutworm, Apamea devastator (native in New 

York and Ohio; Volger and Stressler 2011). The latter is an indiscriminate herbivore known to 

feed on C. palustre and may help control Marsh Thistle; however, this moth feeds on a broad 

spectrum of additional plants.

Larvae of the artichoke plume moth (Platyptilia carduidactyla) also feed on Marsh Thistle, but 

as its common name suggests, this species is considered a pest to artichokes. Furthermore, the 

moth’s native range is south of the Great Lakes (Winston et al. 2008). Occasionally Cheilosia 

corydon, a fly native to Italy, feeds on Marsh Thistle (Winston et al. 2008). This fly was released 

in Oregon in 1991 to control several invasive thistle populations. However, since its release, C. 

corydon populations have attacked native and exotic thistles indiscriminately (ODA Plant 

Division 2011). Additional insects that feed on and/or use C. palustre for part of their life cycle 

are listed on these websites (Lindsey 2005, Hogan n.d.).

Goats are attracted to the flowering stage of many thistles, including C. palustre. Only about 

0.5% of thistle seeds that pass through their digestive systems remain viable, making it unlikely 

that they would aid in the spread of this species. Effective grazing could reduce Marsh Thistle

populations, although it is unclear whether grazing would ultimately control C. palustre via the 

trampling of rosettes or facilitate its spread through the creation of safe sites for germination 

(Fraser 2000). Reseeding of native vegetation may enhance the success of prior control efforts. 

Moreover, goats do not select for Marsh Thistle and may also eat native thistles in intermingled 

communities (Popay and Field 1996).

Van Leeuwen found that a combination of European grazers (rabbits, the hoverfly Cheilosia 

grossa, and Epiblema scutula) resulted in an approximately 30% reduction in flower heads on 

Cirsium palustre. Furthermore, plants that had suffered predation had a reduced stem height, 
222

resulting in a reduced seed dispersal distance of surviving achenes (van Leeuwen 1983). 

Additional research is needed to determine if native species of rabbits and insects could have 

similar results on controlling C. palustre in the Great Lakes.

Physical

All physical control efforts need to be carefully executed and monitored for several years before 

reducing C. palustre infestations (GLIFWC 2006, Sheehan 2007).

Where infestations are small, hand pulling may be effective. In Chequamegon-Nicolet National 

Forest (CNNF), Wisconsin, individual plants were mechanically controlled by cutting the root 

just below the surface with a spade (USDA Forest Service 2005a, GLIFWC 2006). This method 

is most effective if completed before flowering so that all plant material can be left on site to 

decompose (Invasive Plant Council of British Columbia 2008). If this method is implemented

while flowers and seeds are present, flower heads must be bagged and removed from site; the 

remaining plant material can be left on site (Invasive Plant Council of British Columbia 2008).

Mowing before plants flower may reduce the release of seeds (OLA and MAFF 2002); however, 

there is a risk of regrowth with extra flower heads, and extensive mowing of rosettes could 

promote growth of this early successional species once mowing ceases (Fraser 2000, Nordin et 

al. 2008). Repeated close mowing can reduce a C. palustre infestation in three to four years 

(Gumbart 2012.). Mowing a minimum of three times per growing season can be enough to 

weaken the following year’s population (Boos et al. 2010). However, in a study by Falinska 

(1999), the density of C. palustre seeds in the soil decreased dramatically as time since last 

mowing increased, indicating that frequent mowing may actually increase the density of Marsh 

Thistle in the seedbank.

Little is known about Marsh Thistle response to fire as a control strategy (Gucker 2009).

Chemical

Foliar spray with clopyralid (e.g., Transline®, Curtail®, Reclaim®) or metsulfuron-methyl is the 

preferred chemical treatment for C. palustre in CNNF, Wisconsin (USDA Forest Service 2005b). 

However, glyphosate (e.g., Rodeo®, Aquamaster®, Roundup®) must be used in areas that are 

wet or near open water (USDA Forest Service 2005b). Either of these two treatments are most 

effective if applied in the spring when plants are 6 to 10 inches tall and still in the budding stage 

or applied directly to the flower heads in the fall (USDA Forest Service 2005b, Boos et al. 2010). 

To minimize damaging other non-target species when using glyphosate, stems should be cut 

close to the ground and a small amount sprayed onto the cut area (Sheehan 2007).

Conium maculatum

Regulations (pertaining to the Great Lakes region)

Conium maculatum is a well-established invasive species in Ohio and has been designated as a 

prohibited noxious weed (Ohio Division of Natural Areas and Preserves and Nature Conservancy 

2000, USDA NRCS 2012). The possession, transportation, transfer, or introduction of Poison 

Hemlock is restricted in all Wisconsin counties except: Crawford, Dane, Grant, Green, Iowa, 

Lafayette, Richland, Rock, and Sauk (WIDNR 2010a). The Robert W. Freckman Herbarium of 
223

the University of Wisconsin lists an eradication notice for C. maculatum (Robert W. Freckman 

Herbarium 2012). However, as of 2007 it was not illegal to sell Poison Hemlock in Wisconsin 

(Annen 2007).

Regionally, the Great Lakes Indian Fish & Wildlife Commission (GLIFWC) classify this species 

as capable of causing moderate to severe ecological impacts and/or having limited effective 

control options available (Falck and Garske 2003).

Control

Physical or chemical removal of C. maculatum individuals is relatively easy, but complete 

eradication may be difficult due a viable seed bank and reintroductions. Management efforts may 

need to be continued for several years to be effective (J. McHenry pers. comm. in Pitcher 1985).

Biological

The Poison Hemlock defoliating moth (Agonopterix alstroemeriana) was accidentally introduced 

to the United States, but it is now being investigated as a potential biocontrol agent because of its 

monophagous (feeding on a single food source) association with C. maculatum (Castells and 

Berenbaum 2006). Hemlock Moth larvae feed on the young stem tissue, flowers, and seeds 

(Forest Health Staff 2006d). High densities of A. alstroemeriana have been effective drivers of 

plant mortality in C. maculatum stands in the western United States, where several hundred 

larvae have been reported from a single plant. However, as a chemical defense, alkaloid 

production appears to increase with A. alstromeriana herbivory, potentially driving surviving 

populations to higher levels of toxicity over time (Castells et al. 2005). Furthermore, A. 

alstroemeriana was found to be targeted by a predatory wasp (Euodynerus foraminatus) in 

Illinois, suggesting that the effectiveness of biocontrol may be lessened in the Midwest and other 

locations where E. foraminatus exerts top-down pressure on A. alstroemeriana (Castells and 

Berenbaum 2008). Although A. alstroemeriana is widespread in the United States, larvae may 

still be difficult to obtain for biocontrol purposes (Castells and Berenbaum 2006).

Trichoplusia ni, the cabbage looper, is a generalist lepidopteran that is found throughout the 

United States and occasionally feeds on C. maculatum. Overall growth of T. ni is not stunted, but 

larvae raised on diets enriched with the piperidines found in C. maculatum develop slower. A 

prolonged larval stage makes T. ni more vulnerable to predators and could reduce overall 

biocontrol capabilities (Castells and Berenbaum 2008).

Papulio poluxenes, Black Swallowtail Butterfly, will lay eggs on C. maculatum, but a study 

conducted in central New York found low larvae survivorship (Feeny et al. 1985).

Conium maculatum is capable of being infected by multiple viruses, including Ring Spot Virus, 

Carrot Thin Leaf Thin Virus (CTLV), Alfalfa Mosaic Virus (AMV), and Celery Mosaic Virus 

(CeMV) (Howell and Mink 1981). However, viral infections appear to stunt growth rather than 

cause mortality, diminishing their potential for biocontrol (Howell and Mink 1981, Pitcher 

2004). Another disadvantage to using these types of biological control agents is the potential for 

them to escape in neighboring habitats, especially agricultural fields (J. McHenry pers. comm. in 

Pitcher 1985).
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Physical

Care should be taken in handling this toxic plant; it is recommended that gloves are worn 

(Pitcher 2004). If any body part comes into contact with any part of the plant, be sure to wash it 

thoroughly (OLA and MAFF 2002). Hand pulling or digging out the taproot are effective method 

of control for small populations, especially when the soil is moist (WIDNR 2008). Mowing close 

to the ground is another option of mechanical control if the blade is close to the ground (WIDNR 

2008). A dust mask should be worn for protection to avoid inhaling toxins while mowing (King 

County 2011). In both cases, efforts are most effective if completed before the plants flower and 

multiple follow-up efforts should be taken to prevent regrowth or new growth (Parsons 1973, 

Pitcher 2004, WIDNR 2008, Woodard 2008). Poison Hemlock remains for several years after 

death and should be removed where there is a risk of consumption by livestock, wildlife, or 

children (Pitcher 2004).

Chemical

Effective control of large infestations may require chemical agents (WIDNR 2008, King County 

2011). Chemical control of rosettes (before flowering) is a common form of management for 

Conium maculatum because it is such a prolific seed producer (Forest Health Staff 2006d, 

Woodard 2008). Application of herbicides early in spring when sprouts are just emerging may 

also result in effective control, but if C. maculatum has a large presence in the seed bank, 

multiple applications may be needed (Forest Health Staff 2006d).

Several herbicides—including chlorsulfuron, hexazinone, imazapic, glyphosate (Round-Up®), 

metribuzin, metsulfuron, picloram, triclopyr, terbacil, imazapic plus glyphosate, and metsulfuron 

plus 2,4-D plus dicamba—were found to be the most effective chemical agents (Pitcher 1985, 

Jeffery and Robinson 1990, OLA and MAFF 2002, Forest Health Staff 2006d, Woodward 2008, 

King Country 2011). The surrounding plant community should be surveyed prior to selecting a 

herbicide. The effectiveness of all herbicides declines over time, so multiple applications are

recommended from spring to fall (Woodard 2008).

Extra care should be taken when using herbicides near desired wild vegetation or agricultural 

crops. Foliar herbicides applied with a wick will minimize damage to other nearby plants if C. 

maculatum is growing amongst favorable vegetation (Jeffery and Robinson 1990, OLA and 

MAFF 2002). Glyphosate and metsulfuron is not recommended for use near croplands 

(Monsanto Company 2007, WIDNR 2008). In a study conducted by Jeffery and Robinson 

(1990), hexazinon, metribuzin, and terbacil controlled Poison Hemlock with damaging alfalfa 

when applied while the alfalfa was still dormant. For more in-depth instructions on the use of 

2,4-D, glyphosate, and/or metsulfuron, refer to the Pacific Northwest Weed Management

Handbook (Prather et al. 2014).

Echinochloa crus-galli L. P. Beauvois

Regulations (pertaining to the Great Lakes region)

In Minnesota, E. crus-galli is considered to pose a “minimal” threat to ecosystems: poses 

insignificant competition with native species, may naturalize, alters ecosystems insignificantly, 

and has little possibility of spread within or to other sites (Minnesota Invasive Species Advisory 

Council 2009).
225

In 2003, the Great Lakes Indian Fish & Wildlife Commission (GLIFWC) reported that E. crusgalli has widespread established populations, limited effective control methods, and causes low 

to moderate ecological impacts; as a result, GLIFWC does not require its regulation (Falck and 

Garske 2003).

Control

Biological

The fungal pathogen Exserohilum monoceras has shown some success in controlling Barnyard 

Grass (Catindig et al. 2009).

Physical

Echinochloa crus-galli seeds need to be near the surface to germinate; less than one cm of soil 

will inhibit germination (Adamus et al. 2001). Shallow tillage repeated during the spring can 

reduce emergence of Barnyard Grass (OLA and MAFF 2002). Mowing is unlikely to be 

effective because it will stimulate new growth from lateral buds (OLA and MAFF 2002).

Placing mulch over areas where E. crus-galli is expected to emerge will keep the soil cool and 

help suppress germination (Cornell University 2012). Further control methods may be needed.

Chemical

There is a high amount of genetic variation among E. crus-galli communities. Managers who 

choose a chemical control method may need to adjust the compound/application rate to fit the 

needs of the site (Altop and Mennan 2011). Mature plants show little sensitivity to herbicides 

applications. Herbicides applied pre-emergence or shortly after emergence typically exhibit the 

most effective control (Ahmadi et al. 1980, Maun and Barrett 1986, OLA and MAFF 2002).

Barnyard Grass is susceptible to sulfometuron methyl (Oust XP®, Spyder®), clethodium (Select 

MAX®, Intensity®), glyphosate (Accord®, Foresters’ Glypro®, Roundup®, Cornerstone®, 

Razor®), imazapyr (Aresenal AC®, Habitat®, Chopper®), linuron (Linex 4L®, Loroz DF®), 

norflurazon (Predict®), sethoxydim (Sethoxydim E-Pro®, Poast Plus®). It is also susceptible to 

simazine (Simazine 4L®, Simazine 90 DF®), fluazifop (Fusilade DX®), hexainon (Velpar®), 

pendimethalin (Pendulum 3.3 EC®); however it should be noted that these herbicides cannot be 

used in Forest Sustainability Certified Areas (Keely and Thullen 1991, WIDNR 2011).

Ahmadi et al. (1980) found that when applied to Barnyard Grass five cm in height, glyphosate, 

terbuthryn, paraquat, atrazine and buthidazole all resulted in 100% control. The efficacy of 

herbicides containing glyphosate was increased when applied when soil was moist, because it 

allowed for better translocation through the entire plant (Ahmadi et al. 1980). Glyphosate plus 

2,4-D will also control E. crus-galli; it is most effective when applied six days after the last 

irrigation or rainfall (Wicks and Hanson 1995).

Haloxyfop, fluazifop, and sethoxydim offer effective control of (Balyan and Malik 1989). 

Seedling growth of E. crus-galli can be effectively reduced by applying fluazifop and haloxyfop 

(separately) to the soil (Kells et al. 1986).
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Ammonium salt of imazapic (Plateau®) is effective at controlling Barnyard Grass (BASF 

Corporation 2011).

Diclofop was effective at controlling Barnyard Grass, especially when applied postemergence to 

plants with less than four leaves (West et al. 1980).

2,4-D sodium (Hormicide®) applied pre-emergence will help prevent barnyard from growing. 

Young plants can be controlled with paraquat, where as more mature plants can be controlled 

with 2.2-DPA. In Australia E. crus-galli plants treated with F-34 (3,4 - dichloropropionanilide) 

2-3 weeks post-emergence were successfully controlled (FAO 2012).

Bispyribac is a postemergent herbicide that is registered for grasses in rice fields and offers 

effective control of E. crus-galli. Its efficacy may be increased when used in combination with a 

spray adjuvant and/or urea ammonium nitrate (Koger et al. 2007).

Postemergent treatment with propanil and pendimenthalin offers good control of Barnyard Grass

(Setre Chemical Company 1986). Propanil is most effective against Barnyard Grass applied 

before the plants have three leaves (Snipes and Street 1987). However, Barnyard Grass can 

become resistant to propanil and resistant communities have been reported in Arkansas (Gealy et 

al. 2003).

DPX-79406 (1:1 premix of nicosulfuron and rimsulfuron) and rimsulfuron are registered for use 

in Ontario and offer good control of Barnyard Grass growing intermingled with corn (Bosnic and 

Swanton 1997). When applied post-emergence, Rimsulfuron plus thifensulforn resulted in 97% 

control (Krausz et al. 2000).

Cyhalofop-butyl offers good control of E. crus-galli when applied early postemergence. When 

applied to Barnyard Grass growing within rice, this compound had only slight effects on rice 

quantity and quality (Ntanos et al. 2000). The ethyl ester of fenoxaprop will control E. crus-galli 

and most varieties of rice are tolerant (Snipes and Street 1987).

For more specific information on chemical control methods, please visit:

Cornell University’s Pesticide Management Education Program, Pacific Northwest Weed 

Management Handbook, and The Rice Knowledge Bank.

Other

Ongoing research indicates that E. crus-galli may be susceptible to various natural, 

biodegradable herbicides derived from microorganisms and from other plant species (Malik 

1997, Khanh et al. 2006, Kato-Nogucki et al. 2012, Li et al. 2012a).

Epilobium hirsutum L.

Regulations (pertaining to the Great Lakes region)

Great Hairy Willow Herb is ranked as having “moderate environmental invasiveness” by the 

New York State Office of Invasive Species and is considered “well-established” by the Ohio 

Department of Natural Resources (Ohio Division of Natural Areas and Preserves and Nature 
227

Conservancy 2000, NYISC 2010). The transportation, translocation, or introduction of E. 

hirsutum is prohibited in Wisconsin, except in Kenosha County (Burea of Plant Industry 2012). 

Epilobium hirsutum is listed as an introduced species in Ontario (Canadensys 2012). Regionally, 

the Great Lakes Indian Fish & Wildlife Commission (GLIFWC) classifies this species as capable 

of causing moderate to severe ecological impacts and/or having limited effective control options 

available.

Control

Given the invasive nature of E. hirsutum, control methods need to be applied and monitored for 

several years to be effective (King County 2008). For localized infestations, the GLIFWC 

recommends E. hirsutum “be controlled immediately upon detection before it becomes 

established and spreads” (Falck and Garske 2003).

Biological

Elephant moth (Deilephila elpenor) feeds on Epilobium, but is not a native to the Great Lakes 

(Hoskins 2012, Pittaway 2012). Genetic material extracted from E. hirsutum individuals 

displaying phyllody of flowers and/or plant yellowing revealed infection by epilobium phyllody 

(EpPh) phytoplasma, an obligate, parasitic bacteria that attach to phloem tissue (Alminaite et al. 

2002). The ability of this phytoplasma to act as a biocontrol agent is still unknown. Additional 

insects that feed on and use E. hirsutum for part of their life cycle are listed on the website of J. 

Lindsey.

Physical

Small populations of E. hirsutum can be hand dug, placed into plastic bags, and disposed of in 

the trash (King County 2008). When hand digging, one should be sure to remove as much of the 

root pieces as possible because rhizomes left in the soil can generate new plants (King County 

2008, Campbell et al. 2010). Mowing or cutting of mature plants will not kill the plant, but 

flowering stems can be cut in late summer or early fall to prevent seed production and dispersal 

(King County 2008).

After 18 weeks in water logged and flooded conditions, Lenseen et al. (2000) found E. hirsutum

populations only achieved 82% and 54%, respectively, of the mean biomass growth as 

populations in drained conditions. Furthermore, it was determined that flooded individuals 

experienced reduced rhizomal growth in terms of numbers, size, and biomass (Lenssen et al. 

2000). Growth of water logged plants was further limited by pruning adventitious roots. This 

procedure reduced the depth of the plant’s primary root system and made individuals more 

susceptible to uprooting by various environmental conditions (flooding, wind, etc.) (Etherington 

1984). Both of these experimental insights suggest that combined water level manipulation and 

root pruning may be beneficial to the control of E. hirsutum.

Due to the regenerative nature of rhizomes, composting plant material off-site is not 

recommended (King County 2008).

Chemical

Epilobium hirsutum populations treated with Patron 170 are typically susceptible to severe injury 

or death. It should be noted that this pesticide is currently prohibited by from use on Forest 
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Stewardship Council (FSC) land (WIDNR 2011). Under dry conditions, 2,4-D will control great 

hairy willow-herb (Evans et al. 2003). In moist or aquatic locations, glyphosate (Rodeo®) will 

stress or kill above-ground portions of the plant, but the root system will remain intact and plants 

will recover (Evans et al. 2003).

Frangula alnus P. Mill.

Regulations (pertaining to the Great Lakes region)

The New York Invasive Species Council assessed F. alnus as having a high risk of causing 

ecological harm and recommended that its use be prohibited (NYISC 2010). This species is 

restricted in Wisconsin; it may not be transported, transferred, or introduced into any ecosystem 

(Bureau of Plant Industry 2012). Frangula alnus is considered an exotic weed by the Illinois

Department of Natural Resources; sale of this species within the state is not allowed (Bureau of 

Environmental Programs 2009). It is listed as a restricted noxious weed in Minnesota and the 

importation, sale, or transport this plant is illegal (MNDNR 2009).

This species is not widespread in the ceded territories governed by the Great Lakes Indian Fish &

Wildlife Commission (GLIFWC). The GLIFWC recommends that Glossy Buckthorn be 

controlled immediately upon being found. In areas where F. alnus is already present, the 

GLIFWC categorizes it as capable of having severe ecological impacts and recommends that 

small, peripheral populations be controlled upon detection and center populations be monitored 

(Falck and Garske 2003).

Control

When treating a large infestation and/or working with limited resources, priority should be given 

to the largest trees bearing blooms or fruits (Thompson and Luthin 2004). It is important to have 

a disposal method in place for the portions of the Glossy Buckthorn that contain fruit. Stems and 

branches with berries can be destroyed by burning; those without fruit can be left on site to 

decompose (USDA NRCS 2007). If burning is not an option, fruit should be disposed of off-site 

(PADCNR n.d.).

Biological

Currently, there are no specific biological control agents for this species, but research on more 

generalized herbivores is ongoing (Chandler et al. 2010).

Physical

Cutting alone will not control this species because it will resprout, regardless of what time of 

year it was cut (Brock 2012). When using a physical control method, effort should be made to 

limit soil disturbance so as not to cause to erosion (Larson 2009).

Individual plants less than 0.5 inch in diameter can be removed by hand. Removing manually is 

easiest when the soil is sandy or is moist (MNFI 2012). Care must be taken to disturb the soil as 

little as possible (Buenzow 2010). Plants that are 0.5 inch to 1.5 inch in diameter can be 

physically removed with a mechanical device- care should be taken to disturb as little of the soil 

as possible (Buenzow 2010).

Glossy Buckthorn can also be controlled through girdling. In this method, cuts are made to the 

trunk or main stem just above the base, the bark is removed (including the green, cambium layer 
229

beneath the bark) (USDA NRCS 2007). The cut should be large enough, about an inch long, to

prevent the tree from healing. The Illinois Natural History Survey recommends making two

parallel cut 4-5 inches apart when girdling (Heidorn 2011). This method is most effective when 

in the summer after the leaves have fully developed or after the leaves have dropped off in the 

early winter (USDA NRCS 2007). This method is less effective on plants that have many main 

trunks/main stems.

If Glossy Buckthorn is growing in a grassland or savanna ecosystem, controlled burns may offer 

long-term control. This method needs to be repeated ever 2-3 years (MNDNR 2009). Burns are 

most effective from April through June and from September through November (Hanson et al. 

2012).

Repeated mowing in open areas has been reported to be effective in prevented seedling 

establishment (Ohio EPA 2001). This method is most effective for plants less than two years old. 

Mowing in early spring and again in fall will help deplete the energy reserves in the root system, 

deplete the seed bank, and will not interfere with any birds that may use Glossy Buckthorn for 

nesting (USDA NRCS 2007).

If Glossy Buckthorn is present in a managed wetland with a lowered water level, returning the 

level to its original depth may flood and kill the plants. The impact a changed water level will 

have on the whole ecosystem should be determined in advance (Roman 2007).

Chemical

Adding dye to herbicides prior to application, will help distinguishes between plants that have 

and have not been treated (MNFI 2012).

One method is to spray foliage with herbicides such as glyphosate (Accord®, Foresters’ 

Glypro®, Roundup®, Cornerstone®, Razor®), triclopyr (Garlon 3A®, Garlon 4®, Tahoe 4E®), 

fluazifop (Fusilade II®), imazapyr, metsulfuron-methyl, 2,4-D, or picloaram to control Glossy 

Buckthorn (WIDNR 2011, Hanson et al. 2012). Glyphosate will kill any vegetation it comes in 

contact with and triclopyr will kill broadleaf plants, but will not harm grass if applied properly 

(MNDNR 2009). The best time to use the foliage spray method is between May and November 

(Hanson et al. 2012).

For plants with stems less than six inches in diameter, basal steam treatment, in which an oilbased herbicide is applied directly the bark from the root collar up 12-18 inches, can be used 

without having to cut down the plant (MNDNR 2009, Buenzow 2010). Glyphosate and triclopyr 

can also be used for this technique (Hanson et al. 2012). Herbicide can be applied with a lowpressure backpack sprayer. Herbicide can be applied any time of year, providing there is access 

to the ground line; although late fall and winter are preferred because plants are dormant. Glossy 

Buckthorn may leaf out in the spring after a fall or winter herbicide application, but the leaves 

should senesce as the chemicals are translocated throughout the plant (Buenzow 2010). Basal 

spraying is the most cost-effective method of controlling populations of Glossy Buckthorn

(Thompson and Luthin 2004).
230

Both the ester and amine formulations of triclopyr can be used to treat Glossy Buckthorn; the 

amine form is safe for use in most wetlands (Larson 2009, MNFI 2012).

Other

Larger trees can be cut near soil level in late summer or early fall. The stumps should then be 

treated within two hours of being cut with herbicides containing triclopyr. Only the cut surface 

needs to be treat when using a water-soluble herbicide, but when using an oil-based herbicide, 

treat the cut surface and the remaining bark (MNDNR 2009).

Glyceria maxima

Regulations (pertaining to the Great Lakes region)

Glyceria maxima poses a high ecological threat to ecosystems; therefore, the New York Invasive 

Species Council recommends that this species be prohibited (NYISC 2010). The New Invaders 

Watch Program lists G. maxima on its “watch list” for Illinois (Maurer 2009). G. maxima is 

prohibited from transport, transfer, or introduction in Wisconsin; however, there are exceptions 

made for several counties (Bureua of Plant Industry 2012).

The Great Lakes Indian Fish & Wildlife Commission (GLIFWC) has not detected G. maxima 

within its territories. To keep it from invading, the GLIFWC recommends controlling any 

individuals of this species immediately (Falck and Garske 2003).

Control

G. maxima is a perennial species; therefore, populations may require treatment for two to three 

years for complete control (USACE 2012b).

Biological

There are no known biological control methods for this species.

Physical

Small infestations of G. maxima can be dug up; care should be taken to remove all parts of the 

roots and rhizomes (Forest Health Staff 2006e). Subsequent removal of seedlings germinated 

from the seed bank or missed rhizomes pieces may be necessary (King Country 2012). Small, 

dense communities of Reed Mannagrass can also be controlled by being covered with black 

plastic for five or six weeks during the growing season (Forest Health Staff 2006e).

The vegetative spread of larger populations can be controlled by repeated mowing, cutting, 

harvesting, roto-tilling, or rotovating (Sundblad and Robertson 1988, USACE USACE 2012b) 

Where applicable, these treatment methods can be supplemented with artificially created flood 

conditions (Hroudová and Zákravský 1999). Mowing or cutting two to three times a summer 

may deplete the energy reserves in the roots and rhizomes. This may reduce G. maxima’s ability 

to compete and allow other vegetation to expand into the site (King County 2012).

Chemical

A foliar spray of glyphosate (3% solution) applied early to late summer will control populations 

of G. maxima (King County 2012, USACE 2012b). Rhizomes may survive after initial spraying 
231

(USACE 2012b). Braverman (1996) found that glyphosate at two kg ai/ha and dalapon (2,2 

dichloropropanoic acid) at 10 kg ai/ha controlled G. maxima. Imazapyr is most effective on Reed 

Mannagrass when applied in summer or early fall and when water levels are low and plant stems 

are not submerged (King County 2012, USACE 2012b).

In floating fens in the Netherlands, sulfate was experimentally added to the soil. This caused the 

free sulfide concentration to increase and resulted in a decrease in the growth of G. maxima 

(Loeb et al. 2007).

Other

For large populations, herbicide treatment will be an effective option. If the decaying plant 

material falls into a nearby body of water and decomposes, the dissolved oxygen levels could 

decrease. To avoid this, dead plant material should be removed two to four weeks after 

herbicides have been applied (King County 2012).

Hydrocharis morsus-ranae L.

Regulations (pertaining to the Great Lakes region)

Prohibited in Illinois, Michigan, Minnesota, and Wisconsin (GLPANS 2008). In Minnesota ̧ 

possession, import, purchase, transport, or introduction of H. morsus-ranae will result in a 

misdemeanor (MN DNR 2013b). 

The New York Invasive Species Council ranks this species as posing a very high ecological 

threat and recommends that it be regulated (NYISC 2010).

The Great Lakes Indian Fish & Wildlife Commission have no found H. morsus-ranae in their 

ceded territories, but recommend immediate control upon detection (Falck and Garske 2003).

Control

Biological

Grass Carp, Ctenopharyngodon idella, feeds on H. morsus-ranae. However, this introduction of 

this species may also have a negative effect on native vegetation, which might outweigh the 

benefits of H. morsus-ranae control (Mikulyuk and Nault 2011). 

Physical

Mowing does not control H. morsus-ranae population (Sager and Clerc 2006). Removing 

manual harvesting may provide temporary control (IL DNR 2009a, WIDNR 2012c). To improve 

efficacy of this method, harvesting should occur in the spring; after the turions have begun 

growing, but before dense mats form (Catling et al. 2003).

Another possible method for small water bodies would be to have a water draw-down after 

turions have germinated, but before extensive growth occurs (Catling et al. 2003).

Chemical

Diquat, imazapyr, penoxsulam, and imazamox offer excellent control of H. morsus-ranae 

(AERF 2013).
232

Impatiens glandulifera

Regulations (pertaining to the Great Lakes region)

Impatiens glandulifera is ranked as having “low environmental invasiveness” by the New York 

State Office of Invasive Species and is therefore unregulated in that state. Regionally, the Great 

Lakes Indian Fish & Wildlife Commission (GLIFWC) classify this species as capable of causing 

moderate to severe ecological impacts and/or having limited effective control options available 

(Falck and Garske 2003).

Control

Regardless of method, all control methods should be adaptive and involving monitoring of the 

site for several years to ensure plants do not germinate from the seed bank (King County 2010a).

It should also be noted that in experiments conducted by Wadsworth et al. (2000), I. glandulifera

was so prolific that even scenarios with 99% control efficiency were as ineffective as scenarios 

with no management action.

Biological

To date, no specific biological control methods are available for I. glandulifera (Sheppard et al. 

2006). However, allowing cattle or sheep access to areas infested with I. glandulifera will 

control the population and the spread of the species either by direct grazing or by trampling of 

young seedlings (CEH 2004b).

The species Aphis fabae, Impatientinum balsamines, and Deilephila elpenor are known to feed 

on Ornamental Jewelweed, but their capacity to act as biological control agents is still unknown 

(Beerling and Perrins 1993). Although an initial experiment by Tanner (2011) indicated that 

Deilephila elpenor exhibited lower biomass and survivorship when raised on I. glandulifera.

In its native range, I. glandulifera has been known to harbor Puccinia komarovii (a rust 

pathogen) which is currently undergoing research as a control agent (Tanner 2011).

Physical

A good control method for small infestations is the removal of Ornamental Jewelweed by pulling 

or digging. Efforts should be concentrated prior to seed-set in the spring, while the soil is still 

moist (King County 2010a).

Hartmann et al. (1995) found that mowing, mulching or soil cultivation were successful in 

controlling I. glandulifera populations in Germany. Mowing infestations of I. glandulifera also 

causes less soil erosion than hand-pulling or digging (King County 2010a). Ornamental 

Jewelweed should be cut close to the ground, preferable below the lowest node to prevent 

regrowth (CEH 2004b). If the vegetative parts are to be left on-site to decompose, plant material 

should be allowed to dry out completely or the stems should be crushed (by walking or jumping 

on them) to prevent regrowth. Flower heads and seed capsules should not be left on site (King 

County 2010a).
233

Chemical

Herbicides are more effective on populations of I. glandulifera that are large or inaccessible to 

other equipment (King County 2007a). Application of any herbicide needs to be carefully timed. 

Seedlings need to be large enough that they will be covered by the herbicide, but before flowers 

are produced (King County 2010a). Glyphosate (Round-up® or AquamasterTM) is most 

effective if applied to growing leaves. Care should be taken not to get the herbicide on desirable 

plants because it is non-selective and will damage any foliage it comes in contact with (King 

County 2007a). Herbicides containing triclopyr (Renovate3), 2,4-D, or metsulfuron are more 

selective and will not harm most grass species (King County 2007a). These herbicides may be 

preferable if I. glandulifera infestations are in mixed communities and/or near water bodies 

(Centre for Plant Aquatic Management 2004).

When using herbicides, do not cut or mow treated plants until they have died completely, which 

may take two weeks (King County 2007a).

Iris pseudacorus

Regulations (pertaining to the Great Lakes region)

The New York Invasive Species Council ranked Iris pseudacorus as posing a high ecological 

risk and recommends that it be prohibited (NYISC 2010). The Ohio Department of Natural 

Resources lists Yellow Iris as a “well-established invasive” (Ohio Division of Natural Areas and 

Preserves and Nature Conservancy 2000). Under the Michigan Public Acts 74-80 of 2005, I. 

pseudacorus is a prohibited aquatic plant species: a person cannot have any purebred or hybrid 

variant of this species, or fragments or seed unless they are being collected for identification, 

and/or the person is in the process of legally removing/eradicating the species (Latimore et al. 

2011). In Illinois, I. pseudacorus cannot be possessed, propagated, bought, sold, bartered, 

transported, transferred, or loaned with a permit (GLPANS 2008). I. pseudacorus is established 

and considered a “moderate threat” to local ecosystems in Minnesota. This has led it to be 

classified as a restricted species that cannot be planted/released with a permit (GLPANS 2008, 

Minnesota Invasive Species Advisory Council 2009).

In 2001, the Great Lakes Indian Fish & Wildlife Commission (GLIFWC) considered I. 

pseudacorus capable of severe ecological impacts even though it tends to occur in a few small 

populations and has a wide array of control options (Falck and Garske 2003). Yellow Iris was 

still listed as a “high priority” invasive species in 2009, 2010, and 2011 (Falck et al. 2010, 2011, 

and 2012).

Control

For large infestations, it is best to start in the areas with the lowest concentration of Yellow Irises 

and progressing towards high density areas (i.e. periphery toward the center, upstream toward 

downward) (King County 2009).

The glycosides in I. pseudacorus leaves and rhizomes cause skin irritation, so care should be 

taken to protect skin from contact regardless of the control method (Cooper and Johnson 1984, 

Forest Health Staff 2006f, Nature Conservancy n.d.).
234

Biological

Insects and animals do not exert grazing pressure on I. pseudacorus in its native range (Forest 

Health Staff 2006f). Although numerous pathogens and insects attack Yellow Iris, there are 

currently no known biological control agents (Tu 2003).

Physical

Small infestations of Iris pseudacorus seedlings can be easily pulled or dug up by hand, 

especially in damp or wet soil (King County 2009). This method is also feasible for small stands 

of mature plants; however, tools (pickaxes, saws, etc.) may be needed to remove the rhizomes 

(King County 2009). Care should be taken to remove all parts of the rhizomes to prevent 

resprouting (Forest Health Staff 2006f). For populations of Yellow Iris growing in standing 

water, removal of the leaves and stems above water before flowering can result in good control 

and reduced spread (Simon 2008 in Noxious Weed Control Program 2009). Sites should be 

monitored for the emergence of new plants from the seed bank or from rhizome sections that 

were not removed after control measures are completed (King County2009). This control method 

may need to be repeated for three or four year to be effective.

Repeat mowing or removal of seed pods can control the spread of larger infestations (Forest 

Health Staff 2006f). Repeated removal of the aboveground portions of Yellow Iris may also 

deplete the plant’s energy reserves and may eventually kill it (Tu 2003).

Mechanical removal in sensitive areas, such as shallow streambeds, can be expected to cause 

extensive disturbance to the substrate and permit the establishment of other unwanted plants 

(Jacono 2001). Plant material should not be composted on-site because rhizomes can continue to 

growth for up to three months without water (Sutherland 1990).

Burning is not recommended for control because of this plant’s strong tendency to resprout from 

rhizomes (Clark et al. 1998, Sutherland 1990).

Chemical

Applications with herbicides such a glyphosate (RodeoTM or AquamasterTM) or imazapyr 

(HabitatTM) can provide control of larger infestations (Forest Health Staff 2006f, King County 

2009, MN DNR 2012a). If Yellow Iris is mixed with desirable plants species, targeted control 

can involve cutting the stems of I. pseudacorus and applying the herbicide directly to the cut area 

(IISCTC 2007). Areas should not be mowed for several weeks after herbicide application to 

allow the treatment to be effective (King County 2009).
235

Other

Cutting followed by herbicide (glyphosate) treatment with a dripless wick may be the best 

method for controlling plants in sensitive sites.

Juncus compressus Jacq.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Aphids may occasionally feed on Juncus spp., but most rushes are fairly resilient to extensive 

damage from insect or diseases (Stevens and Hoag 2003). Cattle, horses, and sheep graze on 

Juncus spp., but the extent of control gained from grazing is unknown (CEH 2004a, Cosyns et al. 

2005).

Physical

There are no known physical control methods for this species.

The rhizome matrix of Juncus spp. enables it to withstand periods of drought and flooding; water 

level fluctuation is not recommended as a physical control method (Stevens and Hoag 2003).

Chemical

Sethoxydim (Vantage®) will target most grass species and should not affect nearby broadleaf 

herbs, sedges, or woody plants (Brock 2012). Glyphosate and ammonium salt of imazapyr 

(Plateau®) will control J. compressus, but are non-selective and should be applied carefully 

(CEH 2004a) Glyphosate should be sprayed directly onto foliage in mid-late summer (CEH 

2004a).

Juncus gerardii Loisel.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Juncus gerardii seedlings that emerge in late winter or early spring are taller and fertile and can 

be distinguished from the seedlings that emerge at the end of spring or beginning of summer and 

are shorter and infertile (Bouzillé et al. 1997).
236

Biological

Aphids may occasionally feed on Juncus spp., but most rushes are fairly resilient to extensive 

damage from insect or diseases (Stevens and Hoag 2003). Cattle, horses, and sheep graze on 

Juncus spp., but the extent of control gained from grazing is unknown (CEH 2004a, Cosyns et al. 

2005).

Physical

There are no known physical control methods for this species.

The rhizome matrix of Juncus spp. enables it to withstand periods of drought and flooding; water 

level fluctuation is not recommended as a physical control method (Stevens and Hoag 2003).

Efforts made to graze an area with J. gerardii may increase the size of its seed bank (Jutila 1998, 

Jutila 1999).

Chemical

Sethoxydim (Vantage®) will target most grass species and should not affect nearby broadleaf 

herbs, sedges, or woody plants (Brock 2012). Glyphosate and ammonium salt of imazapyr 

(Plateau®) will control J. gerardii, but are non-selective and should be applied carefully (CEH 

2004a). Glyphosate should be sprayed directly onto foliage in mid-late summer (CEH 2004a).

Other

Charpentier et al. (1998) found that a water depth of 10 cm and salinity level of 2 g/L NaCl 

caused J. gerardii to stop shoot and root growth and decrease seed production. A constant water 

level and a low salinity level may limit the growth and expansion of J. gerardii populations 

(Charpentier et al. 1998).

Juncus inflexus L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Aphids may occasionally feed on Juncus spp., but most rushes are fairly resilient to extensive 

damage from insect or diseases (Stevens and Hoag 2003). Cattle, horses, and sheep graze on 

Juncus spp., but the extent of control gained from grazing is unknown (Centre for Aquatic Plant 

Management 2004a, Cosyns et al. 2005).

Physical

Juncus inflexus will regrow if it is cut or mowed; therefore, these methods will only provide 

control if done repeatedly (Missouri Botanical Garden 2012).

The rhizome matrix of Juncus spp. enables them to withstand periods of drought and flooding; 

water level fluctuation is not recommended as a physical control method (Stevens and Hoag 

2003).
237

Chemical

Sethoxydim (Vantage®) will target most grass species and should not affect nearby broadleaf 

herbs, sedges, or woody plants (Brock 2012). Glyphosate and ammonium salt of imazapyr 

(Plateau®) will control J. inflexus, but are non-selective and should be applied carefully (CEH 

2004a). Glyphosate should be sprayed directly onto foliage in mid-late summer (CEH 2004a).

Lupinus polyphyllus

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Lupines usually require ecological disturbance to persist. Control is generally unnecessary in 

undisturbed sites.

Biological 

There are no known biological control methods for this species. Lupines may be toxic and 

populations often increase in grazed (pasture) systems. Several native insects feed on Lupines, 

but are considered insufficient for control (DiTomaso 2013). 

Physical

Hand pulling, tillage, and digging are effective for controlling established plants, but the 

disturbance from these methods can promote new recruitment. The root system should be 

severed below the thickened crown. Mowing is not effective unless done frequently enough to 

prevent seed production. Fire is not an effect control as this promotes germination (DiTomaso 

2013). 

Chemical

Several alternatives are available for chemical control – most are most effective when applied 

post-emergence and before flowering. 2,4-D and/or dicambia can be applied at temperatures less 

than 26.6oC. Glyphospate is effective for spot treatment where reseeding (with natives) is 

planned as it will not injure seedlings. Chlorsulfuron and metsulfuron are also effective 

(DiTomaso 2013).

Lycopus asper

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods specific to this species.

Lycopus europaeus L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
238

Control

There are no known biological, physical or chemical control methods specific to this species.

Lysimachia nummularia

Regulations (pertaining to the Great Lakes region)

New York Invasive Species Council determined that this species has a very high ecological 

impact and recommends that this species be prohibited within the state (NYISC 2010).

Control

Biological

There are no known biological control methods for this species.

Physical

Small populations can be pulled up by hand; all plant fragments should be removed to prevent 

resprouting (Kennay and Fell 2011, MISIN and MNFI 2013d, PADCNR n.d.).

A prescribed burning in the spring, when L. nummularia is green and native plants are still 

dormant, can be an effective means of control (Kennay and Fell 2011). This control method may 

need to be repeated for several years for complete control (Tu et al. 2001).

Given the growth structure of Moneywort, mowing will not control this species (Kennay and Fell 

2011, PADCNR n.d.). Planting native grasses after any physical control method could shade out 

any potential regrowth from Moneywort (MISIN and MNFI 2013d, PADCNR n.d.).

Chemical 

Rodeo® or Rodeo® will be effective at controlling L. nummularia (Kennay and Fell 2011, 

PADCNR n.d.)

Lysimachia vulgaris

Regulations (pertaining to the Great Lakes region)

The New York Invasive Species Council determined that this species poses a high ecological 

threat and recommends that it be prohibited within the state (NYISC 2010). 

The Great Lakes Indian Fish & Wildlife Commission considers this species capable of causing 

severe ecological impacts and recommends it be controlled within their ceded territories (Falck 

and Garske 2003).

Control

Biological

There are no known biological control agents for this species (King County 2010b).
239

Physical

Hand-pulling is effective for small infestations and/or for young plants (King County 2010b, 

MISIN and MNFI 2013c). Care should be taken to dig out and remove all plant fragments, 

especially the rhizomes, to prevent regrowth (King County 2010b). All seed heads and rhizomes 

pieces should be disposed of in plastic bags and removed from the site (King County 2010b).

Repeated mowing of Lysimachia vulgaris may contain the existing population, but it will not 

eradicate it (King County 2010b).

Chemical 

To control larger infestations, treatment with herbicides containing glyphosate (Rodeo®, 

AquaMaster®), imazapyr (Habitat®), or triclopyr (Garlon 3A®, Renovate 3®) may be necessary 

(King County 2010b). It is important to note that glyphosate and imazapyr are non-selective and 

will harm any plant it comes in contact with (King County 2010b). Tricolpyr will not harm 

grasses, sedges, or cattails, and may be more appropriate to use in diverse plant communities 

(King County 2010b). Physical control methods should not be used on populations treated with 

herbicides until several weeks after application (King County 2010b).

Lythrum salicaria

Regulations (pertaining to the Great Lakes region)

Lythrum salicaria is listed as an exotic weed in Illinois (525 Illinois Compiled Statuses § 10/3

and 10/4) making it illegal to buy, sell or distribute plants, its seeds, or any part without a permit. 

Planting, sale, or other distribution without a permit is also prohibited in Indiana (312 IN 

Administrative Code § 14-24-12). Purple Loosestrife – including all cultivars – is a prohibited 

invasive species in Minnesota (MN Administrative Rules § 6216.0250). The species is restricted 

in Michigan, with an exemption for sterile cultivars (MI NREPA 451 § Section 324.41301). 

Planting or sale of the species without a permit is prohibited in Ohio (Ohio Revised Code §

927.682), though the director may exempt varieties ‘demonstrated not to be a threat to the 

environment’. Pennsylvania has designated all nonnative Lythrum species and their cultivars as 

noxious weeds (7 PA Code § 110.1). Purple Loosestrife is designated both as a restricted species 

(Wisconsin Statutes NR § 40.05) and as an invasive aquatic plant (Wisconsin NR 109.07 (2)) in 

Wisconsin. 

Control

Biological

Biological control agents do not eliminate the target weed, but when successful, can sup- press 

weed populations to a nonsignificant level (Rees et al. 1996). Five species of beetles have been 

approved for the biocontrol of L. salicaria (Blossey et al. 1994ab). 

Galerucella calmariensis and G. pusilla are both leaf-feeding chrysomelids. These beetles 

defoliate and attack the terminal bud area, drastically reducing seed production. The mortality 

rate to Purple Loosestrife seedlings is high. Evidence of Galerucella spp. damage are round 

holes in the leaves. Four to six eggs are laid on the stems, axils, or leaf underside. The larvae 

feed constantly on the leaf underside, leaving only the thin cuticle layer on the top of the leaf. 

Initial introductions in eastern North America occurred in Virginia, Maryland, Pennsylvania, 
240

New York, Minnesota, and southern Ontario in August 1992 (Hight et al. 1995). In 1992, these 

three beetles were released in Washington. Galerucella spp. populations visibly impacted Purple 

Loosestrife stands by 1996 (Washington State Department of Ecology 2012). In the Great Lakes 

region, Sea Grant conducted an extensive, multi-state program involving youth in raising and 

releasing Galerucella beetles for control of Purple Loosestrife (Michigan Sea Grant 2001).

Hylobius transversovittatus is a root-mining weevil that also eats leaves. This beetle eats from 

the leaf margins, working inward. The female crawls to the lower 2-3 inches of the stem then 

bores a hole to the pithy area of the stem where 1-3 eggs are laid daily from July to September. 

Or, the female will dig through the soil to the root, and lay eggs in the soil near the root. The 

larvae then work their way to the root. H. transversovittatus damage is done when xylem and 

phloem tissue are severed, and the carbohydrate reserves in the root are depleted. Plant size is 

greatly reduced because of these depleted energy reserves in the root. The larvae evidence is the 

zig-zag patterns in the root. 

Nanophyes marmoratus and N. brevis are seed eating beetles. Young adults feed on new leaves 

on shoot tips, later feeding on the flowers and closed flower buds. Sixty to one hundred eggs are 

laid in the immature flower bud. Seed production is reduced by 60%. There were two test sites 

releases in 1996. Approval to introduce N. marmoratus was granted followed by introductions in 

New York and Minnesota in 1994. Additional releases occurred in New Jersey in 1996. N. 

marmoratus has also been released in Ohio (Ohio EPA 2001). Release of N. brevis planned for 

1994 was delayed due to contamination of the original shipment with a parasitic nematode 

(Piper, 1997). This infection appeared benign for N. brevis; however, due to the potential for 

non-target effects of the nematode after introduction into North America, only disease free 

specimens should be introduced, which, at present, effectively precludes the introduction of N. 

brevis (Blossey 2002a).

Bayeriola salicariae, a gall midge, was studied and screened between 1990 and 1992 (Blossey 

and Schroeder 1995). Based on results indicating a potential wider host range, the gall midge B. 

salicariae was not proposed for introduction (Blossey and Schroeder 1995).

Targetted grazing by sheep has also been used as a biocontrol (Kleppel and LaBarge 2011).

Revegetation of disturbed riparian sites can be used to prevent Purple Loosestrife establishment 

and to reduce re-establishment after control procedures are applied. Fowl Mannagrass (Glyceria 

striata), Foxtail Sedge (Carex alopecoidea), and Reed Canarygrass (Phalaris arundinacea) have 

achieved dominance and prevented re-invasion in plots where Purple Loosestrife was 

experimentally removed. Smartweed (Polygonum lapathifolium) is reported to out-compete 

Purple Loosestrife during its first year of growth. Seeding Japanese Millet (Echinocloa 

frumentacea, also called billion-dollar grass) at 30 pounds/acre on exposed moist soil after 

drawdown and before Purple Loosestrife seedlings began to grow provided control. Japanese 

Millet is considered an exceptional wildlife plant (Jacobs 2008).

Physical

Most mechanical and cultural attempts to control Purple Loosestrife are ineffective. A

single known exception is cutting followed by flooding.
241

For small infestations and isolated plants, hand pulling may be effective. Pull individual 

loosestrife plants by hand before seed is set. The entire root system must be removed, but do not 

dig out roots because soil disturbance may release seeds buried in the soil and break off plant 

parts, which then reproduce. Instead, a cultivator may be used to tease roots from the soil. All 

plant parts should be bagged to prevent dispersal or resprouting and preferably burned. Followup treatments are recommended for at least three years.

Frequent cutting of the stems at ground level is effective but must be continued for several years

(Courtney 1997). Mowing is generally not effective as it exposes the seed bank. 

Flooding is generally ineffective at controlling Purple Loosestrife, though some success has been 

reported for control of seedlings when using flooding regimes in excess 30 cm for over seven

weeks (Balogh 1986). 

Prescribed burning is not an effective management tool for Purple Loosestrife. The dead upright

stems do not carry fire well and the fine fuels are often lacking. The growing points of the root

crown are about two cm (0.8 inch) below the soil surface, so surface fires are not likely to inflict

much damage. Purple Loosestrife begins spring growth about a week or 10 days after 

broadleaved cattails, so a fire of sufficient intensity to damage Purple Loosestrife could also 

damage desirable native species (IL DNR 2009a).

Chemical 

Only herbicides permitted for wetland use may be used to control Purple Loosestrife. There are 

four chemicals that can be used to manage Purple Loosestrife on sites with standing or moving 

water typical of where it invades. Triclopyr and glyphosate are used most commonly. However, 

2,4-D, and imazapyr are also formulated for aquatic applications. Widespread elimination of 

standing biomass may result in the exposure and sprouting of the immense Purple Loosestrife

seed bank. Thus, broadleaf-specific herbicides which do not harm monocot species (such as 

common wetland grasses and sedges) are preferred. The most species specific way to apply 

herbicide is by cutting and treating the stems. Foliar spray can be used by applying herbicide 

after the period of peak bloom, in late August. Any control method should be followed up on a 

yearly basis to catch any missed plants or new sprouts (Ohio EPA 2001)

Marsilea quadrifolia

Regulations (pertaining to the Great Lakes region)

Marsilea quadrifolia is a prohibited plant species in in Illinois (GLPANS 2008). 

The Great Lakes Indian Fish & Wildlife Commission have not found M. quadrifolia within their 

ceded territories, but recommend that it should be controlled immediately if found (Falck and 

Garske 2003). 

Control

Biological

There are no known biological control methods for this species.
242

Physical

There are no known physical control methods for this species.

Chemical 

In studies conducted in Japan, M. quadrifolia was susceptible to the herbicide bensulfuron 

methyl (Aida et al. 2004, Luo and Ikeda 2007).

Mentha aquatica L.

Regulations (pertaining to the Great Lakes region)

The Great Lakes Indian Fish & Wildlife Commission considers this species capable of causing 

severe ecological impacts and recommends it be controlled within their ceded territories (Falck 

and Garske 2003).

Control

Biological

Chrysolina herbacea feeds on M. aquatic, despite the deterrents this species produces to 

minimize damage caused by herbivores (Atsbaha Zebelo et al. 2011).

Physical

Hand-pulling may control small populations of Mentha spp. (MISIN and MNFI 2013d).

Chemical 

General herbicides, such as glyphosate, are effective at controlling Mentha spp. (MISIN and 

MNFI 2013d).

Mentha gracilis Sole (pro sp.)

Regulations (pertaining to the Great Lakes region)

In 2011, the Great Lakes Indian Fish & Wildlife Commission ranked Mentha gracilis to be a 

lower priority for regulation and management (Falck et al. 2012).

Control

Biological

Production of Gingermint can be significantly reduced by diseases such as rhizome rot 

(Rhizoctonia solani Kuhn.) (Skotland 1979, Skotland and Traquair 1982), septoria leaf spot 

(Septoria menthae Oudem.) (Green 1961) and verticillium wilt (Verticillium dahliae Kleb.), and 

insects such as mint flea beetle (Longitarsus waterhousei Kutschera) and mint bud mite 

(Tarsonemus spp.). Further, Mentha gracilis is susceptible to both races of mint rusts that affect 

its parent mint species (Johnson et al. 1999). However, none of these have not been evaluated for 

biocontrol of invasive populations (Poovaiah et al. 2006). 

Physical

Hand-pulling may control small populations of Mentha spp. (MISIN and MNFI 2013d).
243

Chemical 

General herbicides, such as glyphosate, are effective at controlling Mentha spp. (MISIN and 

MNFI 2013d).

Mentha spicata

Regulations (pertaining to the Great Lakes region)

In 2011, the Great Lakes Indian Fish & Wildlife Commission ranked M. spicata to be a lower 

priority for regulation and management (Falck et al. 2012).

Control

Biological

Several diseases of mints are well characterized as crop pests – e.g., Mint Rusts – and activity of 

these pests appears to be quite host specific (Johnson 2013) but the potential for use of mint 

diseases for biocontrol has not been the subject of direct research.

Physical

Hand-pulling may control small populations of Mentha spp. (MISIN and MNFI 2013d).

Soil barriers may be used to restrain rhizomatous spread if plants are grown in borders or other 

areas where spread is unwanted. Removal of flower spikes will stimulate new vegetative growth 

(Missouri Botanical Garden 2013).

Chemical 

General herbicides, such as glyphosate, are effective at controlling Mentha spp. (MISIN and 

MNFI 2013d).

Myosotis scorpioides L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. Wisconsin has a proposal to list this species as 

restricted.

Control

Control options have not been very well documented. This species is likely very difficult to 

control dueto abundant seed production and spread via stolons.

Biological 

There are no known biological control methods for this species. 

Physical

This plant cannot survive exposure to temperatures below -33oF (USDA NCRS 2012).

Myosoton aquaticum (L.) Moench

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species. 
244

Control

Control options have not been very well documented. 

Biological 

There are no known biological control methods for this species. 

Physical

Mow before seeds form, reseed heavily infested areas with competitive forage species (Doll et al.

2004). 

Chemical

Dicamba or glyphosate are considered effective controls (Doll et al. 2004).

Myriophyllum spicatum L.

Regulations (pertaining to the Great Lakes region)

Myriophyllum spicatum is a prohibited species in Illinois and Michigan; its hybrids and variants 

are also prohibited in Minnesota and Wisconsin (GLPANS 2008). In Michigan, a person cannot 

knowingly possess a live organism (Latimore et al. 2011). In Minnesota, it is illegal to possess, 

import, purchase, sell, propagate, transport or introduce Eurasian Watermilfoil (Falck et al.

2011).

The Great Lakes Indian Fish & Wildlife Commission listed this species as a “high priority” for 

control within their ceded territories (Falck et al. 2012).

Control

Due to decades of university, state and federal research and experience with M. spicatum in the 

United States and Canada, several methods have been developed to help in its management.

The best way to minimize the spread of M. spicatum is to remove any visible plant fragments and 

rinse all equipment; allow them to dry completely before using them in another waterbody (IL 

DNR 2009a). 

Biological

Since 1963, the Grass Carp, Ctenopharyngodon idella has been released to suppress Eurasian 

Watermilfoil and other nuisance aquatic plants in numerous sites within North America (Julien 

and Griffiths 1998, CEH 2004d). It has been found that Grass Carp may only eat Eurasian 

Watermilfoil after native plants have been consumed (IL DNR 2009a). To achieve control of 

Eurasian Watermilfoil generally means the total removal of more palatable native aquatic species 

before the Grass Carp will consume Eurasian Watermilfoil. In situations where Eurasian 

Watermilfoil is the only aquatic plant species in the lake, this may be acceptable. However, 

generally Grass Carp are not recommended for Eurasian Watermilfoil control (Washington State 

Department of Ecology 2013).

Laboratory research has shown that the fungus Mycoleptodiscus terrestris reduces the biomass of 

M. spicatum significantly and may be a possible biocontrol (biopesticide) agent (IL DNR 2009a).
245

A North American weevil, Euhrychiopsis lecotie, may be associated with natural declines at 

northern lakes (Sheldon 1994, Creed and Sheldon 1995). Euhrychiopsis lecotei feeds on the new 

growth of M. spicatum and can help keep populations under control; it is common for the 

populations of for E. lecotei and M. spicatum to exhibit the classic predator-prey cycles (Creed 

and Sheldon 1995, Michigan Sea Grant 2012b). Studies have found the herbivorous weevil to 

cause significant damage to Eurasian water-milfoil while having little impact on native species, 

suggesting the insect as a potential biocontrol agent (Creed and Sheldon 1995). Female weevils 

have great fecundity when raised the on M. spicatum as opposed to native M. sibiricum (Solarz 

and Newman 1996, Creed 1998, TNC Vermont 1998, Sheldon and Jones 2001).

Physical

Because this plant spreads readily through fragmentation, mechanical controls such as cutting, 

harvesting, and rotovation (underwater rototilling) should be used only when the extent of the 

infestation is such that all available niches have been filled. Using mechanical controls while the 

plant is still invading will tend to enhance its rate of spread.

Mechanical harvesting has been widely used in the Midwest (RICRMC 2007). Small populations 

of Eurasian Watermilfoil, such as those around docks or in swimming areas, can be removed by 

hand-pulling and/or the use of a sturdy handrake (Bargeron et al. 2003) Multiple harvests within 

the same growing season will yield the best results (Bargeron et al. 2003). If multiple harvests 

are not possible, the single harvest should happen before peak biomass, in early summer, 

otherwise regrowth will occur (Bargeron et al. 2003, WI DNR 2012b). Large equipment exists to 

mechanically remove milfoil in larger areas (Bargeron et al. 2003). Dredging is also effective 

method of removal (CEH 2004d). Care should be taken to remove all fragments to prevent 

regrowth or deoxygenation from plant decomposition (CEH 2004d, MISIN and MNFI 2013b). 

Plant fragments can be disposed of by burning, burying, composting (away from the water), or 

by trash disposal (IL DNR 2009a). In Okanagan Lake, British Columbia, authorities have 

apparently successfully experimented with management by simultaneously rototilling plants and 

roots and underwater vacuuming (Newroth 1988).

Where possible, Eurasian Watermilfoil can be drowned or dehydrated by water level 

manipulation (Bates et al. 1985, Bargeron et al. 2003, WIDNR 2012b). Water drawdowns are 

most effective when the plants are exposed to several weeks of drying time and root crowns are 

exposed to sub-freezing temperatures (IL DNR 2009a). This method could have serious effects 

on other aquatic life (IL DNR 2009a).

Water level manipulation is often used conjunction with herbicides and/or shade barriers 

(Swearingen et al. 2002, Bargeron et al. 2003). Localized control (in swimming areas and around 

docks) can be achieved by covering the sediment with an opaque fabric which blocks light from 

the plants (bottom barriers or screens).

Myriophyllum spicatum is also susceptible to ultrasound pulses and this could prove to be a more 

selective physical method of control (USACE 2012b).
246

Chemical 

Numerous chemicals will have an effect on M. spicatum: amine salts of Endothall (Hyrothol 

191®), and Dipotassium Salts of Endothall (Aquathol K®), Diquat dibromide (Reward®), 

Komeen®, cooper, and confentrazone (Water Bureau 2005, RICRMC 2007).

Myriophyllum spicatum is also susceptible to terbutryn (Clarosan 1FG®) and dichlobenil 

(Midstream GSR®, Casoron G® and Luxan®). Treatment with either chemical is most effective 

when applied in spring before the plant is fully grown and in still water (CEH 2004). Herbicides 

containing dichlobenil will also affects rooted submerged plants and some rushes (CEH 2004d).

The amine formulations of 2,4-D granules (Navigate®, Aquakleen®, Aquacide®) are effective 

on controlling Eurasian Watermilfoil and will not damage grasses (Lembi 2003, Water Bureau 

2005, IL DNR 2009a). The liquid formulation of 2,4-D can be used in ponds and lakes at

concentrations less than 2.0 parts per million (Bargeron et al. 2003). This herbicide method is not 

appropriate for large unmanageable areas of milfoil (Bargeron et al. 2003).

One lose-dose application (10 μg/ L) of fluridone (Sonar® and Avast!®) applied in the early 

stages of growth can result in season long control of Eurasian Watermilfoil (Water Bureau 

2005,USACE 2012b, WIDNR 2012b). This application rate resulted in >93% control for a year 

post-treatment in seven out of eight test lakes in Michigan (USACE 2012b). However, the 

Minnesota Department of Natural Resources found that the application rate of 10 parts per 

billions would cause unavoidable damage to native vegetation (Welling 2013).

Liquid triclopyr (Renovate 3® and Renovate® OTF) will provide effective control of Eurasian 

Watermilfoil and is safe to use around grasses and cattails (Lembi 2003, IL DNR 2009a). A 

concentration of 0.75 parts per million of triclopyr was used to control Eurasian Watermilfoil in 

Loon Lake, New York (Miller 2013). 

Light attenuating dyes (Aquashade® and Admiral®) also exhibit effective control of this species

(USACE 2011b).

Najas marina L. 

Regulations (pertaining to the Great Lakes)

Listed as a “species of special concern” in Minnesota; meaning it is extremely uncommon and 

deserves careful monitoring of its status (MN DNR 2013b).

Control

Biological

While waterfowl and fish may consume parts of N. marina; there are no known biological 

control methods for this species (Agami and Waisel 1986, Tarver et al. 1986, Agami and Waisel 

1988).

Physical

There are no known physical control methods for this species.
247

Chemical 

Herbicides containing Endothall (Aquathol K© Liquid, Aquathol Super K ©granular), Diquat 

(Reward©), or Fluridone (Sonar©, Avast!©) are effective against N. marina (Lembi 2003).

Najas minor All.

Regulations (pertaining to the Great Lakes region)

Najas minor is prohibited in Minnesota, Wisconsin, and Illinois (GLPANS 2008). In Minnesota 

it is illegal to possess, import, purchase, sell, propagate, transport, or introduce N. minor or any 

related varieties or hybrids (Invasive Species Program 2011). The New York Invasive Species 

Council ranks this species moderate ecological risk and recommends that the species be 

regulated (NYISC 2010).

The Great Lakes Indian Fish & Wildlife Commission ranked this species as a “low priority” for 

control in 2011, because it was not detected in their ceded territories (Falck et al. 2012).

Control

Biological

There are no known biological control methods for this species (Ohio EPA 2001).

Physical

Manual removal may provide short-term relief by reducing the biomass of N. najas, however, 

small plant fragments may break off and create new plants/infestations (Ohio EPA 2001, 

Robinson 2004, Office of Water Resources 2010).

Benthic barriers (which restrict light and upward growth of submerged plants) may be effective 

in controlling N. minor in high traffic areas: boating lanes, docks, and swimming beaches 

(Robinson 2004). These structures need to be anchored to the sediment and regularly maintained, 

which may impact other benthic and/or plant organisms (Robinson 2004).

Chemical 

Herbicides may be most effective for controlling large populations of Brittle Waternymph

(Office of Water Resources 2010). Herbicides containing amine salts of Endothall (Hydrothol 

191®), dipotassium salt of Endothall (Aquathol K® Liquid, Aquathol Super K® granular), 

Diquat dibromide (Reward®), or Fluridone (Sonar®, Avast!®) will control N. minor (Robinson 

2004, Water Bureau 2005). Cutrine®, Komeen®, Nautique®, and Weedtrine®, will also provide 

effective control of Brittle Waternymph (Robinson 2004, Water Bureau 2005).

Nasturtium officinale

Regulations (pertaining to the Great Lakes region)

Nasturtium officinale is prohibited in Illinois (GLPANS 2008). Even though it is not restricted or 

prohibited, the Wisconsin Department of Natural Resources acknowledges this species as being 

highly invasive and recommends its eradication upon detection (Robert W. Freckman Herbarium 

2012).
248

In 2001, the Great Lakes Indian Fish & Wildlife Commission determined that this species has 

low to moderate ecological impacts, and is difficult to control given the widespread populations 

and/or limited effective control options (Falck and Garske 2003). 

Control

Biological

There are no known biological control methods for this species.

Physical

Manual removal of N. officinale offers control for small populations (WIDNR 2010).

Chemical 

The non-selective compound glyphosate will provide some control; however it will not be 

effective in flowing water and will harm other plant species if it comes in contact (WIDNR 

2010b).

Nymphoides peltata

Regulations (pertaining to the Great Lakes region)

Nymphoides peltata is prohibited in Illinois, Michigan, and Wisconsin (GLPANS 2008). The 

New York Invasive Species Council ranks this species as posing a high ecological risk, and 

recommends that it be prohibited within the state (NYISC 2010).

The Great Lakes Indian Fish & Wildlife Commission has not found N. peltata in their ceded 

territories, but recommend immediate control upon detection (Falck and Garske 2003).

Control

Biological

There are no known biological control methods for this species (CEH 2004c).

Physical

Hand-pulling and mechanical removal is possible because the stems are easily cut by hand tools 

(CEH 2004c, MISIN 2013). Hand raking or using a rope and grapnel is effective when the 

bottom sediments are loose (CEH 2004c). All plants pieces should be removed because new 

plants can grow from broken fragments and/or the decomposing plant material could decrease 

the oxygen levels in the water (CEH 2004c, Kelly and Maguire 2009). Booms or nets can be 

used to catch and remove drifting plant materials (Kelly and Maguire 2009). Even with multiple 

harvests, 100% control is unlikely (CEH 2004c).

Chemical 

Herbicides containing dichlobenil (Midstream GSR®, Casoron G®, Luxan dichlobenil®) are

effective at controlling N. peltata (CEH 2004c). Granules should be sprinkled evenly over this 

species in early spring, after growth has started, but before the leaves reach the water surface 

(CEH 2004c). To ensure eradication, plants may need to be cut back and subjected to a second 

application and/or be treated with another application the following spring (CEH 2004c).
249

Glyphosate can be used to control N. peltata, but it is less effective than dichlobenil and does not 

offer long-term control (CEH 2004c).

Pluchea odorata succulenta (Fern.) Cronq.

Regulations (pertaining to the Great Lakes region)

In Pennsylvania, Pluchea odorata is listed as an endangered species (USDA NRCS 2012).

Control

Biological

The fruit fly Acinia picturata has been known to use P. odorata as a host, but it is unknown if 

this species could be used a biological control agent (Stegmaier 1967).

Physical

There are no known physical control methods for this species.

Chemical 

There are no known chemical control methods for this species

Pluchea odorata odorata (L.) Cass. 

Regulations (pertaining to the Great Lakes region)

In Pennsylvania, P. odorata is listed as an endangered species (USDA NRCS 2012). 

Control

Biological

The fruit fly Acinia picturata has been known to use P. odorata as a host, but it is unknown if 

this species could be used a biological control agent (Stegmaier 1967).

Physical

There are no known physical control methods for this species.

Chemical 

There are no known chemical control methods for this species.

Poa trivialis L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Poa trivialis is used as a turf grass, often overseeded with Bermuda grass. Most research on 

management of this species focuses on promotion of the plants rather than control. Its tendency 

to yellow in the heat of summer makes it undesirable in some turf applications, and most 

information on control is based on these applications.
250

Biological 

Combining herbicides with overseeding an alternative desired grass will help discourage the 

regrowth of surviving P. trivialis and improve overall success of control (Morton and Reicher 

2007).

Physical

There are no known physical control methods for this species.

Chemical

Non selective control with glyphosate (Roundup) followed by reseeding has been the only option 

for control. A herbicide called sulfosulfuron (Certainty®) is currently being labeled by Monsanto 

for selective control of Poa trivialis in Creeping Bentgrass (Street and Sherratt 2013).

Other

This species has low tolerance for high temperatures and drought. 

Polygonum persicaria L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological 

Although P. persicaria plants are susceptible to Arabis Mosaic Virus, no research has been 

undertaken on the development of biological control agents, whether viral or fungal. No 

arthropods known to attack this species specifically have been identified (Plantwise 2013).

Physical

When handling Smartweed, do not place hands close to eyes – it contains a chemical which 

burns.

Management of this species in agricultural settings focuses on elimination of seed as a 

contaminant and cultivation to prevent seedling establishment. Tillage and cultivation which 

disrupts seedlings can be effective. Repeated mowing to prevent seed production can also reduce 

populations over time (Plantwise 2013).

Solarizing black plastic, burning and acetic acid treatments are all effective at killing seeds.

Chemical

P. persicaria seems relatively quick to develop resistance to herbicides – most resources 

discourage use of chemical control or encourage rotation of herbicides with differing modes of 

action (Plantwise 2013). 

Pre-emergent herbicides, especially with the chemical Dichlobenil, are effective. Use on-going 

for up to one year. Systemic herbicides to kill the fibrous roots can also work well. Other options 

for herbicide control are non-selective, contact herbicides. Products containing dicamba work 

best over others containing 2,4-D and glyphosate (Easy Butterfly Garden 2013).
251

Potamogeton crispus

Regulations (pertaining to the Great Lakes region)

Potamogeton crispus is prohibited in Illinois and Minnesota; restricted in Michigan and 

Wisconsin (GLPANS 2008). The New York Invasive Species Council ranks this species as 

posing a “high” ecological risk, and recommends that it be prohibited within the state (NYISC

2010).

As of 2011, the Great Lakes Indian Fish & Wildlife Commission lists this species a high priority 

species and recommends it be controlled within their ceded territories (Falck et al. 2012). 

Control

Biological

The herbivorous Grass Carp, Ctenpharyngodon idella, will provide effective control of P. 

crispus, but may feed on native plants (CEH 2004e). Grass Carp is illegal in some Great Lake 

states (GLIFWC 2006).

Other bottom feeding fish, such as Common Carp, do not feed on P. crispus, but they create 

turbid water conditions and may prevent the growth of this plant species (CEH 2004e).

Physical

Small infestations can be removed manually by cutting, raking, or digging up plants (IISCTC 

2007). The optimal timing for cutting is debated. Some agencies claim that plants should be cut

im early spring and as close to the sediment surface as possible to prevent turion formation 

(MISIN and MNFI 2013a, WIDNR 2012a). Other organizations claim that cutting should not be 

carried out until mid-to later summer to prevent regrowth (CEH 2004e). Regardless when 

cutting/raking occurs, it is important to remove as many plant fragments as possible to limit new 

populations of Curlyleaf Pondweed (PADCNR n.d.). 

When removing this species via digging, root crowns should also be removed from the soil; this 

removal method can be enhanced by the use of a suction apparatus (ENSR International 2005).

The use of equipment such as dredges, underwater rototillers, or hydrorakes are more effective 

for populations in deep waters (ENSR International 2005, USACE 2012b). These physical 

methods are indiscriminate and should only be used on monoculture populations of P. crispus 

(ENSR International 2005). Plant material should be removed after it is cut to prevent regrowth 

or decreases in oxygen concentration due to plant decomposition (ENSR International 2005).

Another option would be to use blankets or other benthic barriers to block sunlight from reaching 

P. crispus (ENSR International 2005). This method will eliminate all vegetation, including 

native species, in 30 – 60 days (ENSR International 2005, GLIFWC 2006). 

In some waterbodies, water draw-down may be an option. All plants, including natives, will be 

exposed to drying or freezing (ENSR International 2005). A water draw-down in autumn may 

kill P. crispus turions and increase the efficacy of this control method (MISIN and MNFI 

2013a).
252

Chemical

P. crispus plants dieback completely in early summer; in order for effective control, herbicides 

should be applied before dieback occurs (MISIN and MNFI 2013a).

P. crispus was effectively controlled by fluridone (Sonar®, Avast!®) in test site lakes in 

Michigan (Getsinger et al. 2001). Control can be obtained with a dose of 6 - 15 ppb with an 

exposure time of 60 – 120 day (ENSR International 2005). This method is only appropriate for 

whole lake applications (IL DNR 2009b).

Endothall (Aquathol K® Liquid, Aquathol Super K® granular) and diquat (Reward®) may offer 

effective control if applied to P. crispus before turion production; typically in April and May 

(ENSR International 2005, WIDNR 2012a). Plants may still continue to grow, but their 

reproductive ability will be greatly reduced (ENSR). Application of either of these chemicals is 

most effective when the water temperature is between 50o– 55o F (IL DNR 2009b). 

Reapplication of diqaut in subsequent years may be necessary for complete control (Bugbee 

2009).

The compounds dichlobenil (Midstream GSR®, Casoron G®, Luxan Dichlobenil® Granules) or 

terbutryn (Clarosan®) will also provide effective control if applied in early spring just as growth 

starts (CEH 2004e)

Herbicides containing 2,4-D will be rapidly taken up by P. crispus, but complete control is 

unlikely (ENSR International 2005).

Puccinellia distans (Jacq.) Parl.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Chemical

Alkali grasses are extremely tolerant of salinity and outcompete other grasses in most brackish 

conditions. Thus management efforts in the freshwater Great Lakes regions targeted at reducing 

salt contamination may benefit efforts to control P. distans.

Rorippa sylvestris (L.) Bess.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Once established, this plant is very difficult to control.

Physical

Readily re-grows from rhizome fragments – most attempts at physical control only exacerbate 

the problem when small fragments are missed and regrow.
253

Chemical

Pre-emergent herbicides suppress top growth but do not kill rhizomes which quickly produce 

new plantlets in response to the loss of the top. Glyphosate is marginally effective. Selective 

post-emergent herbicides are not available (NC State 2012)

Preemergence control was excellent with dichlobenil granules at 3 or 6 lb/A, isozaben at 1 or 2 

lb/A and the geotextile/herbicide (Biobarrier). The geotextile (Typar) fitted as collars alone were 

not effective. Trifluralin incorporated into the surface two inches at 2 lb/A was effective but did 

allow some emergence. Trifluralin plus isoxaben or oryzalin plus isoxaben were also effective at 

rates of 2 plus 0.5, 4 plus 1, or 6 plus 1.5 lb/A, or 3 plus 1, 4.5 plus 1.5, or 6 plus 2 lb/A, 

respectively, of the two herbicide combinations. Metolachlor at 3, 4.5, or 6 lb/A was ineffective 

for preemergence control of three cm rhizome pieces. Post emergence control was not 

commercial with 2,4-D, triclopyr, clopyralid or a combination of the latter two, when treated in 

the 6 to 8 leaf stage with 0.25% or 0.5% solutions (Elmore et al. 1996)

Rumex longifolius DC.

Regulations (pertaining to the Great Lakes region)

There are no federal or state (within the Great Lakes region) regulations for this species.

Control

Biological

Docks are grazed by cattle, sheep, goats, and deer but not by horses. 

Physical

Repeated cultivation is recommended for control of young (seedling) populations. 

Mowing has little effect on established docks, but will prevent seed production. However, 

frequent cutting encourages taproot growth, branching shoots and may aid seedling development 

(from previous year’s seed bank) and so is not recommended. In a pasture heavily infested with 

docks the best option may be to plough and reseed with grass but not immediately. The docks are 

likely to regenerate both vegetatively and from seed, and a period of fallowing or arable cropping 

may help to reduce re-establishment.

Chemical 

Many chemical controls are available for dock species. However, very few are approved for use 

in or near water. Repeated treatments are usually needed to control re-growth.

Dicamba (benzoic acid) is effective on Curly Dock (Rumex crispus) but not on Broadleaf 

Dock (R. obtusifolius). Picloram (pyridine) is effective on most Rumex species. 2,4-DB amine or 

2,4-D ester are effective when applied beforethe flower stalk elongates, but require a 30 day 

withdrawl before feeding as forage. Aminopyralid can be applied to actively growing plants 

before the bud stage. Chlorsulfuron and metsulfuron can be used with young, actively growing 

plants, but should not be used on powdery, dry, or light sandy soils. Sulfoteturon has similar use, 
254

but should not be applied to cropland. Glyphosate can be used at early heading (Pacific 

Northwest Extension 2013).

Rumex obtusifolius

Regulations (pertaining to the Great Lakes region)

The Great Lakes Indian Fish & Wildlife Commission ranked Rumex obtusifolius as capable of 

low to moderate ecological impacts and does not consider it a priority for control within their 

ceded territories (Falcke and Garske 2003). There are no federal (or state within the Great Lakes 

region) regulations for this species.

Control

Biological

Bitter Dock is avoided by rabbits, but it appeared to be a favorite food plant of deer (Amphlett 

and Rea 1909). Docks are grazed by cattle, sheep, and goats, but not by horses. 

Cavers and Harper (1964) list a range of fungi and insects that attack, feed on or occur on docks 

but this not an indication that of their efficacy as control agents. The use of the stem boring 

larvae of the weevils Apion violaceum and A. miniatum for controlling R.obtusifolius has been 

investigated (Hopkins 1980, Freese 1995). In the United Kingdom and elsewhere, there has been 

research on the chrysomelid beetle (Gastrophysa viridula) as a biocontrol agent for both R. 

obtusifolius and R. crispus (Bentley et al. 1980). Larvae of the Leaf-Mining Fly Pegomya 

nigritarsis cause blotch mines on leaves of R. obtusifolius (Whittaker 1994). R. obtusifolius is the 

preferred host plant of Coreus marginatus and has been shown to moderately reduce its seed 

viability (Hruskova et al. 2005). The Leaf Spot Fungus Ramularia rubella causes red spots to 

develop on dock leaves but has no major effect on plant survival. The Rust fungus Uromyces 

rumicis is also non-systemic but has been shown to have some potential as a biological control 

agent (Inman 1971, Schubiger et al. 1986). Dock species are also an alternate host for number of 

viruses, fungus (Dal Bello and Carranza 1995), and nematodes (Townshend and Davidson 1962, 

Edwards and Taylor 1963).

Physical

Caution should be used in physical removal as this plant can cause contact dermatitis.

Repeated cultivation is recommended for control of young (seedling) populations. 

Mowing has little effect on established docks, but will prevent seed production. However, 

frequent cutting encourages taproot growth, branching shoots and may aid seedling development 

(from previous year’s seed bank) and so is not recommended. In a pasture heavily infested with 

docks the best option may be to plough and reseed with grass but not immediately. The docks are 

likely to regenerate both vegetatively and from seed, and a period of fallowing or arable cropping 

may help to reduce re-establishment.

Chemical 

Many chemical controls are available for dock species. However, very few are approved for use 

in or near water. Repeated treatments are usually needed to control re-growth.
255

Dicamba (benzoic acid) is effective on Curly Dock (Rumex crispus) but not on Broadleaf 

Dock (R. obtusifolius). Picloram (pyridine) is effective on most Rumex species. 2,4-DB amine or 

2,4-D ester are effective when applied beforethe flower stalk elongates, but require a 30 day 

withdrawl before feeding as forage. Aminopyralid can be applied to actively growing plants 

before the bud stage. Chlorsulfuron and metsulfuron can be used with young, actively growing 

plants, but should not be used on powdery, dry, or light sandy soils. Sulfoteturon has similar use, 

but should not be applied to cropland. Glyphosate can be used at early heading (Pacific 

Northwest Extension 2013).

Salix alba

Regulations (pertaining to the Great Lakes region)

In 2011, the Great Lakes Indian Fish & Wildlife Commission ranked S. alba to be a low priority 

for regulation and control (Falck et al. 2012). There are no known federal or state (within the 

Great Lakes region) regulations for this species.

Control

Biological

To our knowledge no research into possible biological control organisms for White Willow has 

been attempted in North America or anywhere else. Candidates for insect control are generally 

not sufficiently specific to avoid damage to native willows.

Physical

Like most river trees willows resprout vigorously from cut stumps, and will usually grow back 

into a tree eventually. Repeated cutting of new stump shoots can eventually kill the trees. But 

given their large root systems, cutting would presumably be needed to be done several times per 

growing season for several years in order to starve the roots. Mowing or weed-whipping might 

be useful for seedlings (GLIFWC 2013). Small seedlings can be handpulled, Larger trees may 

require a weed wrench or machinery to remove the root systems. 

Chemical 

Given willows’ tendency to resprout from the roots, multiple applications will likely be needed 

for control.

The nonselective herbicide glyphosate (available commercially as "Roundup" and "Rodeo") is 

commonly used for treating woody invasives such as Crack Willows. Tryclopyr (Garlon 3A or 

equivalent amine formulation) is also effective against broadleaf and woody plants, and has the 

advantage of leaving grasses and sedges intact. Recently Roundup has been shown to be highly 

toxic to both adult frogs and toads and their tadpoles, probably due to the surfactant 

(polyethoxylated tallowamine, or POEA) in this glyphosate formulation (Relyea 2005). Because 

of this and other as yet unknown effects of various herbicide formulations on the environment, 

herbicide should be applied as precisely as possible and only when needed, using only the 

amount needed to get the job done. Any attempt to control Crack Willows or other invasive 

plants in aquatic habitats must be done using Rodeo or other herbicides formulated for use over 
256

water. Permits are required for herbicide application over water in many states, including 

Wisconsin, Michigan and Minnesota (GLIFWC 2013).

Triclopyr amine is best used on young willows (seedlings) that are actively growing. 2,4-D LV 

ester can be applied when leaves are fully developed and growing (amendable to aerial 

application). Metsulfuron is used on fully leafed-out brush. 

Salix fragilis

Regulations (pertaining to the Great Lakes region)

In 2011, the Great Lakes Indian Fish & Wildlife Commission ranked Salix fragilis to be a low 

priority for regulation and control (Falck et al. 2012). There are no known regulations for this 

species.

Control

Biological

To our knowledge no research into possible biological control organisms for Crack Willows has 

been attempted in North America or anywhere else. Candidates for insect control are generally 

not sufficiently specific to avoid damage to native willows.

Physical

Like most river trees willows resprout vigorously from cut stumps, and will usually grow back 

into a tree eventually. Repeated cutting of new stump shoots can eventually kill the trees. But 

given their large root systems, cutting would presumably be needed to be done several times per 

growing season for several years in order to starve the roots. Mowing or weed-whipping might 

be useful for seedlings (GLIFWC 2013). Small seedlings can be handpulled. Larger trees may 

require a weed wrench or machinery to remove the root systems. 

Chemical 

Given willows’ tendency to resprout from the roots, multiple applications will likely be needed 

for control.

The nonselective herbicide glyphosate (available commercially as "Roundup" and "Rodeo") is 

commonly used for treating woody invasives such as Crack Willows. Tryclopyr (Garlon 3A or 

equivalent amine formulation) is also effective against broadleaf and woody plants, and has the 

advantage of leaving grasses and sedges intact. Recently Roundup has been shown to be highly 

toxic to both adult frogs and toads and their tadpoles, probably due to the surfactant 

(polyethoxylated tallowamine, or POEA) in this glyphosate formulation (Relyea 2005). Because 

of this and other as yet unknown effects of various herbicide formulations on the environment, 

herbicide should be applied as precisely as possible and only when needed, using only the 

amount needed to get the job done. Any attempt to control Crack Willows or other invasive 

plants in aquatic habitats must be done using Rodeo or other herbicides formulated for use over 

water. Permits are required for herbicide application over water in many states, including 

Wisconsin, Michigan and Minnesota (GLIFWC 2013).
257

Triclopyr amine is best used on young willows (seedlings) that are actively growing. 2,4-D LV 

ester can be applied when leaves are fully developed and growing (amendable to aerial 

application). Metsulfuron is used on fully leafed-out brush. 

Salix purpurea

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Cultivars of Salix purpurea are often used for erosion control and most species-specific 

management information stems from this use.

Biological

To our knowledge no research into possible biological control organisms for willows has been 

attempted in North America or anywhere else. Candidates for insect control are generally not 

sufficiently specific to avoid damage to native willows.

Gypsy moths can defoliate Purple Willow and Willow Midges can cause significant (though 

rarely fatal) damage. Willow blight is a serious pest to plantings, particularly plants that have 

been damaged by storms. Beaver can have huge impact on this species and are capable of 

eradicating entire stands (USDA NRCS 2002)

Physical

Like most river trees willows resprout vigorously from cut stumps, and will usually grow back 

into a tree eventually. Repeated cutting of new stump shoots can eventually kill the trees. But 

given their large root systems, cutting would presumably be needed to be done several times per 

growing season for several years in order to starve the roots. Mowing or weed-whipping might 

be useful for seedlings (GLIFWC 2013). Small seedlings can be handpulled, Larger trees may 

require a weed wrench or machinery to remove the root systems. 

Chemical 

Given willows’ tendency to resprout from the roots, multiple applications will likely be needed 

for control.

The nonselective herbicide glyphosate (available commercially as "Roundup" and "Rodeo") is 

commonly used for treating woody invasives such as Crack Willows. Tryclopyr (Garlon 3A or 

equivalent amine formulation) is also effective against broadleaf and woody plants, and has the 

advantage of leaving grasses and sedges intact. Recently, Roundup has been shown to be highly 

toxic to both adult frogs and toads and their tadpoles, probably due to the surfactant 

(polyethoxylated tallowamine, or POEA) in this glyphosate formulation (Relyea 2005). Because 

of this and other as yet unknown effects of various herbicide formulations on the environment, 

herbicide should be applied as precisely as possible and only when needed, using only the 

amount needed to get the job done. Any attempt to control Crack Willows or other invasive 

plants in aquatic habitats must be done using Rodeo or other herbicides formulated for use over 

water. Permits are required for herbicide application over water in many states, including 

Wisconsin, Michigan and Minnesota (GLIFWC 2013).
258

Triclopyr amine is best used on young willows (seedlings) that are actively growing. 2,4-D LV 

ester can be applied when leaves are fully developed and growing (amendable to aerial 

application). Metsulfuron is used on fully leafed-out brush. 

Solanum dulcamara

Regulations (pertaining to the Great Lakes region)

After determining that Solanum dulcamara poses a low ecological impact, the Nature 

Conservancy gave this species a “low” national priority ranking (NatureServe 2008). The New 

York Invasive Species Council determined that this species poses a moderate ecological risk, and 

therefore recommended that this species be regulated (NYISC 2010). 

As of 2011, the Great Lakes Indian Fish & Wildlife Commission ranked S. dulcamara as a 

“lower priority” (Falck et al. 2012).

Control

Biological

There are no known biological control methods for this species (King County 2010c).

Physical

S. dulcamara may be controlled by manually digging up the roots; most effective with the plants 

are young and the soil is moist (Waggy 2009). All parts of the roots should be removed to 

prevent regrowth (King County 2010c). Gloves should be worn when handling this species to 

prevent skin irritation (King County 2010c).

Mowing is not a practical method for control because this species can resprout from the 

suckering roots and rhizomes; however it may be useful when manually removing the root 

system (King County 2007b). If mowing is the only option, it must be done several times during 

the growing season to be effective (Forest Health Staff 2006a). Another option would be to cover 

the cut plants with a heavy geotextile cloth for two years to prevent photosynthesis and regrowth 

(King County 2010c). 

Chemical 

Larger infestations may require the use of an herbicide (Forest Health Staff 2006a). The 

herbicide Clopyralid (Transline®, Stinger®, Reclaim®, and Curtail®) is effective on Solanaceae 

(Tu et al. 2001). Triclopyr is effectively taken up in the woody stems, roots, and leaves of 

Bittersweet Nightshade. It is also unlikely to harm nearby grasses, sedges, rushes, cattails, lilies, 

and irises (King County 2010c).

Solidago sempervirens L.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
259

Control

Seaside Goldenrod is being actively planted for dune restoration and wildlife habitat on the east 

coast. Most of the available research-based management information focuses on growing the 

plant rather than control.

Biological 

There are no known biological control methods for this species.

Physical

Spreads only slowly other than by seed, so deadheading or other cutting prior to seed-set can be 

effective for control.

Chemical

This is a salt-adapted species. Efforts to control salt-pollution in the freshwater Great Lakes 

region may help to control populations of this species.

Sparganium glomeratum (Laestad.) L. Neum.

Regulations (pertaining to the Great Lakes region)

Under the synonym, Sparganium erectum, this species is federally listed as a noxious weed 

(Code of Federal Regulations § Title 7, 360.200). It is additionally listed as prohibited in 

Minnesota (MN Administrative Rules § 6216.0250).

Control

Little to no information is readily available on control of S. glomeratum.

Trapa natans

Regulations (pertaining to the Great Lakes region)

Trapa natans is prohibited in Illinois, Michigan, Minnesota, New York, and Wisconsin 

(GLPANS 2008). 

The Great Lakes Indian Fish & Wildlife Commission have not found T. natans in their ceded 

territories, but recommended immediate control upon detection (Falck and Garske 2003).

Control

An integrated management plan that incorporates multiple methods of control will be most 

effective at controlling populations of T. natans. Invaded habitats should be monitored for up to 

12 years after control measures are complete to ensure that the seed bank is exhausted 

(PADCNR n.d., Swearingen et al. 2002).

Biological

In its native range in China, the leaf beetle Galerucella birmanica has significant negative 

impacts on T. natans populations (Ding et al. 2006). However, this species has many other host 

species in the United States, making it unsuitable for use as a biocontrol agent (Maryland Sea 

Grant 2012).
260

Physical

Smaller populations can be controlled by hand harvesting or raking because the roots are easily 

uplifted from the sediment (Naylor 2003). Larger populations, including those thick enough to 

clog waterways, may require the use of a large aquatic plant harvester (PADCNR n.d.). 

Harvesting methods should be conducted before plants set seeds-- typically in July (Maryland 

Sea Grant 2012). All plant fragments, especially those containing roots, should be removed to 

prevent the expansion of the T. natans population (Swearingen et al. 2002). Plant fragments 

should be disposed of far from the water, preferably in a plastic bag (PADCNR n.d.).

260,000 lbs. of Water Chestnut were removed by mechanical means and the help of over 60 

volunteers from the Sassfras River (Maryland) during a three day harvest in 1999 (Naylor 2003). 

Mechanical removal methods have been used annually in Sodus Bay, New York since the 1960s, 

but the T. natans population persists (US EPA 2000). However, mechanical removal followed by 

an application(s) of 2,4-D was able to eradicate a population of T. natans in Maryland (Naylor 

2003).

Laboratory and greenhouse studies by Wu and Wu (2007) demonstrated that ultrasonic waves of 

20 kHz, aimed directly at Water Chestnut stems and petioles, for 10 seconds resulted in 100% 

plant death. 

Chemical 

Herbicides containing 2,4-D (both the amine and butoxy-ethyl ester formulations) have been 

effective in controlling T. natans (USACE 2012b, WIDNR 2012e) Applying 2,4-D just as plants 

are reaching the surface of the water, in early summer, will provide the best results (USACE 

2012b). This compound causes minimal adverse effects on neighboring wildlife (Maryland Sea 

Grant 2012).

Herbicides containing triclopyr are also effective at controlling T. natans, but it is non-selective 

and may harm other plant life (USACE 2012b).

The growth and expansion of Water Chestnut populations can also be repressed if light 

attenuating dyes are applied prior to plant germination (USACE 2012b).

Typha angustifolia L.

Regulations (pertaining to the Great Lakes region)

This species is restricted in Wisconsin; it may not be transported, transferred, or introduced into 

any ecosystem (Bureau of Plant Industry 2012).

Control

Biological

Muskrat (Ondatra zibethicus) populations can have a serious impact on Typha populations; 

however, large populations of muskrats can shift to other plants species and have a long-term 

detrimental effect on the vegetation community (Miklovic 2000).
261

The native boring-moth larvae (Arzama spp.) have been reported to cause damage to Typha

stands, but their use as a species specific biological control is unknown (Miklovic 2000).

Heavy grazing will eliminate Typha spp. from riparian corridors; however, this technique might 

also affect other native species (Stevens and Hoag 2006).

Physical

Mowing during the growing season, once just before the flowers reach maturity and again about 

a month later (when new growth is 2-3 feet high), will kill at least 75% of Narrow-Leaved 

Cattails (Stevens and Hoag 2006).

Burning may also be effective at controlling Typha; however, it needs to be repeated several 

times. Unless the flames have access to the belowground portions of cattails, the rhizomes will 

resprout and grow new plants (Forest Health Staff 2006c, USDA Forest Service 2012). This 

treatment option might also be unfeasible in wet ecosystems or sensitive natural areas (Miklovic 

2000).

Typha spp. are sensitive to the ethanol produced from anaerobic respiration. Flooding a wetland 

could trigger this reaction and help control Typha (Miklovic 2000). Manually digging up plants 

or cutting stems, followed by raising the water level by three inches above the plants will yield 

effective control, as well (Forest Health Staff 2006c).

Chemical

Typha spp. can be controlled by 2,4-D, glyphosate (Rodeo®, Eagre®, AquaNeat®, 

Pondmaster®, Aquapro®, Avocet®, Shore-Klear®, Touchdown Pro®), impazapyr (Arsenal 

AC®, Habitat®, Chopper®, Aquapier®, Gullwing Avocet®), and diquat (Harvester®, 

Redwing®, Reward®, Weedtrine D®) (Water Bureau 2005, Forest Health Staff 2006c, WIDNR 

Division of Forestry 2011, AERF 2012). Glyphosate can result in greater than 80% control 

(Thorsness et al. 1992).

Wick, broom, and/or foliar applications are appropriate techniques for these herbicides (Ohio 

EPA 2001, Forest Health Staff 2006c, Borland et al. 2009). Due to the energy reserves in the 

extensive root system, re-treatments may be necessary (Ohio EPA 2001).

Veronica beccabunga

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Noted by several sources as ‘spreading quickly but easy to control. However, we have not been 

able to track down details for recommended control methods. 
262

A.14 Platyhelminthes

Bothriocephalus acheilognathi Yamaguti, 1934

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Bath treatments are effective control methods for B. acheilognathi infections. Baths should 

contain Droncit® (praziquantel), isopropyl alcohol, and water yielding a final mixture 

concentration ≥ 0.67 ppm praziquantel. Fish densities during treatment should be no greater than 

60 mg fish/L and exposure should last 24 hours. After 24 hours, the treatment should be drained, 

worm parts discarded, and clean water added. After 72 hours, the treatment should be drained 

and worm parts discarded. Fish should then be transferred to a decontaminated container 

(Mitchell and Darwish 2009).

Bothriocephalus acheilognathi infections can be treated with chemically enhanced feed. Drugs 

should be mixed in oil and sprayed on feed at a rate of 1 L/70 kg dry weight. Effective chemicals 

and doses include dibutylin oxide or dibutylin dilurate (250 mg/kg fish) fed over three days 

(Mitchell and Hoffman 1980), Yomesan® (500 g/500 kg dry pellets) fed at 1.5% of body weight 

2–3 times weekly, and Yomesan® (28 g/40 kg) fed for three days (Korting 1974, Mitchell and 

Hoffman 1980, Brandt et al. 1981).

Other

Bothriocephalus acheilognathi populations in aquaculture and ponds can be controlled by 

managing the intermediate host (i.e., copepods) population densities. Effective ectoparasiticides 

include Neguvon®, Masoten®, Dipterex®, Bromex®, and Naled® (Paperna 1996).

Dactylogyrus amphibothrium Wagener or Wegener, 1857

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Implementation of Eurasian Ruffe management may potentially decrease Dactylogyrus

amphibothrium prevalence due to host specificity. However, Ruffe management is considered by 

some (e.g., Ogle 1998) to be difficult and impractical given that the species has developed 
263

several adaptations to compensate for high mortality rates (Lind 1977) and populations rebound 

quickly (Lelek 1987).

Physical

Establishment of quarantines may prevent D. amphibothrium transmission (Reed et al. 1996)

Chemical

D. amphibothrium-specific treatments are unknown. However, multiple chemicals are effective 

at treating monogenean fluke infections in aquaculture systems. Effective benzimidazoles 

include levamisole (Buchmann 1997) and praziquantel, which has high efficacy against 

Dactylogyrus spp. (Buchmann 1997, Schmahl and Mehlhorn 1985). Effective bath treatments 

include formaldehyde (30-100 ppm), sodium chloride, copper sulphate, hydrogen peroxide, 

sodium percarbonate (Buchmann and Kristensen 2003), formalin (25 mg/L for prolonged 

exposure or 150-250 mg/L for 30 minutes), and potassium permanganate (2 mg/L for prolonged 

exposure or 10 mg/L for 30 minutes) (Reed et al. 1996). Effective organophsophate bath 

treatments include metrifonate (0.25-0.5 ppm) and dichlorvos (0.25-0.5 ppm) (Sarig et al. 1965).

Pond infestations can be controlled with formalin (30 mg/L) or trichlorfon (Lepidex®; 0.5 mg/L) 

(Reed et al. 1996). However, monogenean eggs display chemical resilience and therefore the 

above chemical treatments are ineffective at destroying eggs (Reed et al. 1996, Rowland et al. 

2007). Chemical toxicity varies considerably between monogeneans and fish species. Toxicology 

and tolerance tests are suggested prior to using anthelmintics (“dewormers”). Managers are 

encouraged to consider specific host drug tolerance, temperature, salinity, organic material 

content, and drug retention time prior to treatment (Buchmann and Bresciani 2006). Freshwater 

fish species can also be dipped in saltwater to minimize external parasite numbers prior to 

stocking (Reed et al. 1996).

Other

Lampricide TFM may effectively eliminate up to 97% of Ruffe, potential carriers of D. 

amphibothrium, with minimal non-target mortality (Crosier et al. 2012).

Dactylogyrus hemiamphibothrium Ergens, 1956

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

Biological

Implementation of Eurasian Ruffe management may potentially decrease Dactylogyrus

hemiamphibothrium prevalence due to host specificity. However, Ruffe management is 

considered by some (e.g., Ogle 1998) to be difficult and impractical given that the species has 

developed several adaptations to compensate for high mortality rates (Lind 1977) and 

populations rebound quickly (Lelek 1987).
264

Physical

Establishment of quarantines may prevent monogenean fluke transmission (Reed et al. 1996).

Chemical

D. hemiamphibothrium specific treatments are unknown. However, multiple chemicals are 

effective at treating monogenean fluke infections in aquaculture systems. Effective 

benzimidazoles include levamisole (Buchmann 1997) and praziquantel, which has high efficacy 

against Dactylogyrus spp. (Buchmann 1997, Schmahl and Mehlhorn 1985). Effective bath 

treatments include formaldehyde (30-100 ppm), sodium chloride, copper sulphate, hydrogen 

peroxide, sodium percarbonate (Buchmann and Kristensen 2003), formalin (25 mg/L for 

prolonged exposure or 150-250 mg/L for 30 minutes), and potassium permanganate (2 mg/L for 

prolonged exposure or 10 mg/L for 30 minutes) (Reed et al. 1996). Effective organophsophate 

bath treatments include metrifonate (0.25-0.5 ppm) and dichlorvos (0.25-0.5 ppm) (Sarig et al. 

1965).

Pond infestations can be controlled with formalin (30 mg/L) or trichlorfon (Lepidex®; 0.5 mg/L) 

(Reed et al. 1996). However, monogenean eggs display chemical resilience and therefore the 

above chemical treatments are ineffective at destroying eggs (Reed et al. 1996, Rowland et al. 

2007). Chemical toxicity varies considerably between monogeneans and fish species. Toxicology 

and tolerance tests are suggested prior to using anthelmintics (“dewormers”). Managers are 

encouraged to consider specific host drug tolerance, temperature, salinity, organic material 

content, and drug retention time prior to treatment (Buchmann and Bresciani 2006). Freshwater 

fish species can also be dipped in saltwater to minimize external parasite numbers prior to 

stocking (Reed et al. 1996).

Other

Lampricide TFM may effectively eliminate up to 97% of Ruffe, potential carriers of D. 

hemiamphibothrium, with minimal non-target mortality (Crosier et al. 2012).

Dugesia polychroa Schmidt, 1861

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Ichthyocotylurus pileatus

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.
265

Neascus brevicaudatus von Nordmann, 1832

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Scolex pleuronectis Müller, 1788

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Timoniella sp.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

A.15 Protozoa

Acineta nitocrae

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Glugea hertwigi Weissenberg, 1911

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Heterosporis sp.

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.
266

Control

Biological

There are no known biological control methods for this species.

Physical

Effective physical control methods include complete desiccation of holding tanks and equipment 

for 24 hours, freezing at -20°C for 24 hours, and culling (Sutherland et al. 2006, GLFHC 2012b).

Chemical

Immersion of gear in a 2200 ppm bleach (0.7L of bleach per 20 L of water) solution for five

minutes will destroy the parasite (IDNR 2005, Sutherland et al. 2006, GLFHC 2012b).

Other

Infected fish or fish parts should not be discarded back into the water body.

Myxobolus cerebralis Hofer, 1903

Regulations (pertaining to the Great Lakes region)

The Great Lakes Fish Disease Control Policy and Model Program have prohibited stocking the 

Great Lakes and their tributaries with fish from whirling disease infected farms. Fish imported 

into the North Central Region states must be certified free of whirling disease in order to obtain 

import permits (Faisal and Garling 2004).

Ohio requires out of state source facilities to document annual salmonid fish, egg, and sperm 

health inspections for one year prior to importation. Source facilities outside the Great Lakes 

basin must document health inspections for the previous five years with no whirling disease 

occurrences prior to importing salmonids into the Lake Erie watershed (Baird 2005). Indiana

requires source facilities within the Great Lakes basin to document they have been whirling 

disease free for three consecutive years prior to importing salmonid stock. Source facilities 

outside the basin must document salmonid stocks have been whirling disease free consecutively 

since 2002 (Baird 2005). Michigan requires source facilities to document salmonid stocks have 

been whirling disease free for two consecutive years prior to importation, while Wisconsin

requires one. Illinois and Minnesota also require imported salmonid health inspections. 

Minnesota allows the importation of whirling disease infected eggs, if prior egg treatments are 

approved (Baird 2005). Ontario requires an import permit issued by the Canadian Food 

Inspection Agency (CFIA) prior to the importation of certain finfish. Under the Canadian Health 

of Animals Act, aquaculturists are required to report any whirling disease suspicions to the CFIA 

(CFIA 2012).

All eight Great Lakes states (New York, Pennsylvania, Ohio, Michigan, Indiana, Illinois, 

Wisconsin, and Minnesota) have instated similar baitfish regulations to control the spread of 

whirling disease and other fish pathogens. Those of New York include that bait harvested from 

inland waters for personal use is only permitted to be used within the same body of water from 

which it was taken and cannot be transported overland (with the exception of smelt, suckers, 
267

Alewives, and Blueback Herring). Once transported, baitfish cannot be replaced to its original 

body of water (NYSDEC 2012a).

Live or frozen bait harvested from inland New York waters for commercial purposes is only 

permitted to be sold or possessed on the same body of water from which it was taken and cannot 

be transported over land unless under a permit and or accompanied by a fish health certification 

report. Bait that is preserved and packaged by any method other than freezing, such as salting, 

can be sold and used wherever the use of bait fish is legal as long as the package is labeled with 

the name of the packager-processor, the name of the fish species, the quantity of fish packaged, 

and the means of preservation (NYSDEC 2012a).

Certified bait may be sold for retail and transported overland as long as the consumer maintains a 

copy of a sales receipt that contains the name of the selling vendor, date sold, species of fish 

sold, and quantity of fish sold. Bait that has not been certified may still be sold but the consumer 

must maintain a sales receipt containing the body of water where the baitfish was collected and a 

warning that the bait cannot be transported by motor vehicle. Bait sold for resale require a fish 

health certification along with a receipt that contains the name of the selling vendor, date sold, 

species of fish sold, and quantity of fish sold, which must be kept for 30 days or until all bait is 

sold (NYSDEC 2012a).

In addition to baitfish protections, prior to placing fish in New York waters, a fish health 

certification report must document that the fish are whirling disease free.

Control

The following biological, physical and chemical controls only pertain to fish in captive or 

hatchery operations. There are no known control methods of whirling disease in wild 

populations (except for management of spread - see below).

Biological

Managing Tubifex tubifex populations can be implemented as a biocontrol of M. cerebralis. 

Maintaining water quality, reducing favorable habitat by preventing sediment accumulation in 

aquaculture (Crosier et al. 2012), and desiccating holding tanks, equipment, and intake pipes 

may help control T. tubifex (Kaster and Bushnell 1981). Lampricide TFM (3-triflouromethyl-4-

nitrophenol), administered at (4.2-14.0 mg/L) doses, is effective at destroying T. tubifex (Lieffers 

1990). Tubifex tubifex can also be treated in 30°C water for four days, causing triactinomyxon 

(TAM) spore production to stop, thus preventing the next stage of the parasites life cycle (ElMatbouli et al. 1999). T. tubifex ability to support M. cerebralis’ triactinomyxon (TAM) spore 

production may be due to genetic differences among T. tubifex populations. This variability may 

be an important factor in determining infection rates among fish (Baxa et al. 2006) and 

therefore might support certain management practices (Stromberg 2006).

It has been proposed that selective processes are yielding a surviving population of fish that is 

more resistant to M. cerebralis infection on the Madison River, Montana (Vincent 2006). The 

implications of this for management are still unclear. However, research is continuing to 

evaluate the possibility of a developing resistance within salmonid populations (Stromberg 

2006).
268

Physical

Managers have observed that using concrete in aquaculture facilities can reduce the abundance 

of T. tubifex and thus limit the ability of M. cerebralis to reproduce (Mills et al. 1993, Ricciardi 

2001).

The Colorado Division of Wildlife (CDW 2011) administers routine fish health sampling at 

hatchery sites to help slow the spread of M. cerebralis infections by early detection. At the 

Roaring Judy Hatchery, a project is underway to install an ultraviolet system that kills M. 

cerebralis spores (CDW 2011).Treating water with 2537Å UV at doses of 35mWs/cm2can be 

86-100% effective at preventing whirling disease in Rainbow Trout fry (Hoffman 1974) and 

administering 1,300 mWs/cm2of UV under a static collimated beam, can inactivate 100% of the 

TAM spores present (Hedrick et al. 2000).

There is evidence that electricity (1,000 s exposure to low-level DC voltage for 48 hrs) can 

destroy T. tubifex in aquaculture (R. Ingraham and T. Claxton, pers. comm. in Wagner 2002). 

Electrical charges of 1-3 kV pulsed 1-25 times at 99 μsec/pulse is effective at killing large 

numbers of TAM spores (Wagner 2002). Exposing myxospores to 90°C water for 10 minutes is 

also effective at destroying the spores (Hoffman and Markiw 1977).

Experiments by Hoffman (1974) have demonstrated that filtration is not an effective method for 

removing TAM spores from water – due to the small spores size, the filter needed to remove 

them slows flow to rates unacceptable for most applications.

Chemical

Hatchery intake water treated with chlorine (0.5 ppm) administered at two hour intervals once a 

week can reduce infection rates in Rainbow Trout by 63-73% without causing harm to the fish 

(Markiw 1992). Supply water treated with calcium cyanide (488 g/m2) mixed with chlorine gas 

(300 ppm) can be very effective at destroying M. cerebralis spores (Hoffman and Dunbar 1961). 

Water treated with chlorine (130-260 ppm) for 10 minutes may kill 100% of TAM spores present 

(Wagner 2002), and treating with chlorine (5,000 ppm) for 10 minutes is sufficient enough to 

destroy both triactinomyxon and myxospore (E. MacConnell, pers. comm. in Wagner 2002). 

Treating fry with chlorine (10 ppm) for 30 minutes may prevent whirling disease infection 

(Hoffman and O’Grodnick 1977).

It has been demonstrated that feeding Rainbow Trout with pellets containing (0.1%) Fumagillin 

is effective at reducing whirling disease infection. Two groups of Rainbow Trout were 

administered pellets from days 14-64 and 30-160 post infection. Approximately 10-20% of the 

medicated fish harbored spores, whereas 73-100% of non-medicated fish harbored spores (ElMatbouli and Hoffman 1991).

Earthen pond substrate treated with quicklime (CaO) at concentrations >380 g/m2for two weeks 

prior to introducing fish can prevent whirling disease infection by destroying M. cerebralis

spores (Hoffman and Hoffman 1972).

Other

Adherence to local laws regarding transportation of live fish between bodies of water, contacting 
269

local agencies immediately upon noticing signs of whirling disease, properly disposing of fish 

and fish parts, and not transporting mud on boots and shoes between bodies of water are useful in 

controlling the transmission of M. cerebralis spores in the wild.

Psammonobiotus communis

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Psammonobiotus dziwnowi

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Psammonobiotus linearis

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Sphaeromyxa sevastopoli

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.

Trypanosoma acerinae Brumpt, 1906

Regulations (pertaining to the Great Lakes region)

There are no known regulations for this species.

Control

There are no known biological, physical or chemical control methods for this species.
270

A.16 Viruses

Novirhabdovirus sp. genotype IV sublineage b

Regulations (pertaining to the Great Lakes region)

Transportation of Viral Hemorrhagic Septicemia (VHS)-susceptible species out of New York, 

Pennsylvania, Ohio, Michigan, Indiana, Illinois, Wisconsin, Minnesota, Ontario, and Quebec is 

prohibited unless certain conditions are met (USDA APHIS 2008). International movement of 

VHS-susceptible species from the two infected Canadian provinces to the United States is 

permitted if the shipment meets certain requirements and is imported under an APHIS permit for 

direct slaughter, or during catch and release fishing activities (USDA APHIS 2008).

Movement of VHS-susceptible species between VHS-infected or at risk states is permitted as 

long as fish are sent directly to state-inspected slaughter facilities that discharge waste water to a 

municipal sewage system that includes disinfection, or discharge to a non-discharging pond or a 

settling pond that disinfects according to all applicable United States Environmental Protection 

Agency and state regulatory criteria, and are accompanied by a valid VS 1-27 (Permit for 

Movement of Restricted Animals) form issued by an APHIS area office. Remains from slaughter 

facilities must be rendered or composted (USDA APHIS 2008).

Interstate movement of VHS-susceptible fish from VHS-infected or at risk states to non-infected 

states is permitted as long as the fish are accompanied by appropriate state, tribal, or federal 

documentation stating the fish have tested negative for the virus (USDA APHIS 2008). 

Movement of VHS-susceptible species to state, federal, or tribal authorized research and 

diagnostic facilities is also permitted provided that the fish are accompanied by a valid VS 1-27 

form issued by an APHIS area office and the remains are disposed of as medical waste adhering 

to all applicable United States Environmental Protection Agency and state regulatory criteria 

(USDA APHIS 2008).

New York, Pennsylvania, Ohio, Michigan, Indiana, Illinois, Wisconsin, and Minnesota have 

instated similar baitfish regulations to control the spread of VHS and other fish pathogens. Those 

of New York include that bait harvested from inland waters for personal use is only permitted to 

be used within the same body of water from which it was taken and cannot be transported 

overland (with the exception of smelt, suckers, Alewives, and Blueback Herring). Once 

transported, baitfish cannot be replaced to its original body of water (NYSDEC 2012a).

Live or frozen bait harvested from inland New York waters for commercial purposes is only 

permitted to be sold or possessed on the same body of water from which it was taken and cannot 

be transported over land unless under a permit and or accompanied by a fish health certification 

report. Bait that is preserved and packaged by any method other than freezing, such as salting, 

can be sold and used wherever the use of bait fish is legal as long as the package is labeled with 

the name of the packager-processor, the name of the fish species, the quantity of fish packaged, 

and the means of preservation (NYSDEC 2012a).
271

Certified bait may be sold for retail and transported overland as long as the consumer maintains a 

copy of a sales receipt that contains the name of the selling vendor, date sold, species of fish 

sold, and quantity of fish sold. Bait that has not been certified may still be sold but the consumer 

must maintain a sales receipt containing the body of water where the bait fish was collected and 

a warning that the bait cannot be transported by motor vehicle. Bait sold for resale require a fish 

health certification along with a receipt that contains the name of the selling vendor, date sold, 

species of fish sold, and quantity of fish sold, which must be kept for 30 days or until all bait is 

sold (NYSDEC 2012a).

In addition to baitfish protections, prior to placing fish in New York waters, a fish health 

certification report must document that the fish are VHS free.

Ontario has implemented management zones to help slow the spread of VHS. Commercial bait 

operators are prohibited from moving live baitfish out of the VHS Management Zone and live or 

dead bait in or out of the Lake Simcoe Management Zone. Salmon and trout eggs may be 

collected from virus-positive waters only if eggs are disinfected according to the Ministry’s 

protocol. Walleye spawn collection is permitted as long as the fish are stocked back into viruspositive waters. Fish and eggs are permitted to be stocked in waters that are not virus-positive 

only if the facilities are certified VHS free (OMNR 2011).

Control

Biological

There are no known biological control methods for this species.

Physical

Multiple means of control are available to fish hatchery managers, including treatment of water 

with UV light subtype C (280-200nm wavelength) irradiation and heat (>15°C) (McAllister 

1990), exposure to pH levels lower than 2.5 or higher than 12.2, desiccation of tanks and 

equipment (CFSPH 2007), minimization of stressors, cessation of water flow to adjacent 

waterways, and establishment of quarantines (Warren 1983, CFSPH 2007). Furthermore, 

exposure to VHS can be prevented through use of spring water, specific pathogen free (SPF) 

stock, and separate cultivation of salmonids and flatfish (CFSPH 2007).

As with other hitchhiking aquatic species, boaters and anglers are encouraged to clean and 

disinfect their gear (Bakal 2012), as well as to completely drain bilges and live wells before 

moving between bodies of water (OMNR 2012).

Chemical

The VHS virus is sensitive to ether, chloroform, glycerol, formalin, iodophor, sodium hydroxide, 

and sodium hypochlorite, which can be used as disinfectants (CFSPH 2007 McAllister 1990). No 

effective anti-viral agents or commercial vaccines exist (CFSPH 2007). Disinfection of live wells 

and other contaminated equipment can be accomplished with a 10% household bleach/water 

solution (e.g., 100 ml of household bleach to 900 ml of water). Waste water should be discarded 

away from any water body. Virkon® S is another widely available disinfectant (OMNR 2012).
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Other

The United States Fish and Wildlife Service recommend implementation of the International 

Hazard Analysis and Critical Control Point (HACCP) planning standard to prevent the spread of 

VHS (Bakal 2012).

VHS should be reported to Area Veterinarians in Charge (AVIC) or state veterinarians 

immediately upon diagnosis or recognition of the disease. Fish health surveillance programs and 

fallowing are also useful methods of control (CFSPH 2007).

Ranavirus

Regulations (pertaining to the Great Lakes region)

Ohio requires out-of-state source facilities of live fish to provide health inspection and testing 

documentation prior to importation (NCRAC 2010ab). Michigan requires imported aquacultured 

fish to be accompanied by either an official interstate health certificate, official interstate 

certificate of veterinary inspection, or a fish disease inspection report. Importing aquacultured 

fish from source facilities with a record of an emergency disease within the past two years is 

prohibited. Fish imported from non-Michigan source facilities and intended for stocking in 

public waters must be certified free of Largemouth Bass Virus (LMBV) (NCRAC 2010ab). 

Illinois requires source facilities of any species of live fish, eggs, and sperm to document they are 

disease free prior to importation (NCRAC 2010ab). Wisconsin requires source facilities to 

document fish health inspections prior to importing live fish and eggs (NCRAC 2010ab).

Control

Biological

There are no known biological control methods for this species.

Physical

There are no known physical control methods for this species.

Chemical

Disinfection of live wells and other contaminated equipment can be accomplished with a 10% 

household bleach/water solution (i.e., 100 ml of household bleach to 900 ml of water). Waste 

water should be discarded away from any water body. Virkon® S is another widely available 

disinfectant (OMNR 2012).

Other

There is no known cure or effective treatment of LMBV infection (Syska et al. 2012).

Rhabdovirus carpio

Regulations (pertaining to the Great Lakes region)

Importation of live fish, fertilized eggs, and gametes of Spring Viremia of Carp Virus (SVCV)-

susceptible species including Common Carp (Cyprinus carpio), Koi (C. carpio koi), Grass Carp 

(Ctenopharyngodon idella), Silver Carp (Hypophthalamicthys molitrix), Bighead Carp 
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(Aristicthys nobilis), Crucian Carp (Carassius carassius), Goldfish (Carassius auratus), Tench 

(Tinca tinca), Orfe (Leuciscus idus), and Sheatfish (Silurus glanis) is permitted, provided they 

are accompanied by a United States Department of Agriculture import permit and a veterinary 

health certificate (USDA APHIS 2012).

Control

Biological

Single-stranded and double-stranded RNA injections can provide Rhabdovirus carpio protection 

for up to three weeks (Aliken et al. 1996, Masycheva et al. 1995).

Physical

Establishment of quarantines, culling, and stock density reduction during the winter and spring 

are beneficial management practices to prevent the spread of Spring Viremia of Carp Virus 

(SVCV) (CFSPH 2007). R. carpio is inactivated by UV irradiation (254 nm), gamma irradiation 

(103 krads), heating to 60°C for 30 minutes, and exposure to pH 12 for 10 minutes, or pH 3 for 

three hours (CFSPH 2007, OIE 2009).

Chemical

Disinfection of facilities and equipment will prevent the spread of SVCV in aquaculture (CFSPH

2007). R. carpio is susceptible to oxidizing agents like sodium dodecyl sulphate, non-ionic 

detergents, and lipid solvents. The virus is inactivated by formalin (3%) for five minutes, 

chlorine (500 ppm), iodine (0.01%), NaOH (2%) for ten minutes, banzalkonium chloride (100 

ppm for 20 minutes), alkyltoluene (350ppm for 20 minutes), chlorhexidine gluconate (100 ppm 

for 20 minutes), and cresol (200 ppm for 20 minutes) (Ahne and Held 1980, Ahne 1982, Fijan 

1999, CFSPH 2007, Kiryu et al. 2007). Methisoprinol may be useful by inhibiting replication of 

SVCV in vitro. Further testing under culture conditions is necessary (Siwicki et al. 2003).