Ecological Risk Assessment of the Proposed Use of the Herbicide 

Imazapyr to Control Invasive Cordgrass (Spartina spp.) 

in Estuarine Habitat Of Washington State 

Prepared For: 

Washington State Department of Agriculture 

Olympia,WA 98504 

Prepared By: 

ENTRIX, INC. 

Olympia, Washington 

98502 

PROJECT NO. 3000901 

October 30, 2003
Ecological Risk Assessment of the Proposed Use of the Herbicide Imazapyr to 

Control Invasive Cordgrass (Spartina spp.) 

in Estuarine Habitat of Washington State

Prepared For: 

Washington State Department of Agriculture 

1111 Washington Street 

Olympia, WA 98504 

Contract Manager: 

Mr. Kyle Murphy 

(360) 902-1923 

Prepared By: 

ENTRIX, INC. 

148 Rogers St NW 

Olympia, WA 98502 

(360) 352-3225 

Project No. 300901 

Project Manager: 

Jeffrey P. Fisher, Ph.D. 

Contributing Authors: 

J.P. Fisher, B. Mavros, D. Waller, M. Heller, 

B. Suedel, B. Gillespie, J. Slocomb 

October 30, 2003 
TABLE OF CONTENTS 

Page 

1.0 Introduction........................................................................................................................1 

2.0 Problem Formulation ........................................................................................................3 

2.1 Overview of Spartina Infestation in Washington State ...........................................3 

2.1.1 Distribution ..................................................................................................4 

2.1.2 Biology.........................................................................................................7 

2.2 Ecological Receptors, Community Descriptions, and Threatened and Endangered 

Species In Primary Areas Where Spartina is Distributed......................................7 

2.3 Conceptual Model..................................................................................................20 

3.0 Hazard Assessment..........................................................................................................23 

3.1 Mechanism of Action and Efficacy .......................................................................24 

3.1.1 Mechanism of Action.................................................................................24 

3.1.2 Efficacy on Target Vegetation...................................................................25 

3.2 Environmental Fate and Chemistry .......................................................................31 

3.2.1 Physical Chemistry ....................................................................................31 

3.2.2 Environmental Degradation.......................................................................32 

3.3 Toxicity to Terrestrial Receptors...........................................................................40 

3.3.1 Mammals....................................................................................................41 

3.3.2 Birds...........................................................................................................44 

3.3.3 Insects ........................................................................................................45 

3.3.4 Reptiles and Amphibians...........................................................................46 

3.4 Toxicity to Aquatic Receptors...............................................................................46 

3.4.1 Fish.............................................................................................................47 

3.4.2 Aquatic Invertebrates.................................................................................50 

3.4.3 Non-target Aquatic Vegetation..................................................................50 

3.5 Adjuvant and Inert Ingredient Toxicity to Terrestrial and Aquatic 

Ecological Receptors.........................................................................................51 

3.5.1 Inert Ingredients.........................................................................................52 

3.5.2 Surfactants..................................................................................................52 

4.0 Exposure Assessment.......................................................................................................57 

4.1 Estimated Environmental Exposure Concentrations (EEC) ..................................57 

4.1.1 Application Rate ........................................................................................57 

4.1.2 Water Concentrations.................................................................................58 

4.1.3 Expected Plant and Animal Residues........................................................60 

4.1.4 Sediment Concentrations...........................................................................61 

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4.2 Ecological Receptor Exposures .............................................................................62 

4.2.1 Mamalian Exposure ...................................................................................68 

4.2.2 Avian Exposure..........................................................................................71 

4.2.3 Insect Exposure..........................................................................................71 

4.2.4 Reptiles and Amphibian Exposure.............................................................71 

4.2.5 Aquatic Animal Exposure..........................................................................72 

4.3 Adjuvant and Inert Ingredient Exposure Assessment............................................73 

5.0 Risk Characterization to Ecological Receptors from the Use of Imazapyr to Control 

Spartina .............................................................................................................................74

5.0.1 Mammal Risk.............................................................................................74 

5.0.2 Avian Risk .................................................................................................77 

5.0.3 Insects ........................................................................................................78 

5.0.4 Reptiles and Amphibians...........................................................................78 

5.0.5 Fish.............................................................................................................78 

5.0.6 Aquatic Invertebrates.................................................................................79 

5.0.7 Non-target Aquatic Vegetation Risk..........................................................80 

5.1 Relative Hazard and Risk Characterization of Imazapyr, Glyphosate, Surfactants, 

Inert Ingredients, And The Use of No Chemical Control Or Other Agents For 

Spartina Eradication ............................................................................................81 

5.2 Uncertainties and Data Gaps..................................................................................83 

5.3 Conclusions and Recommendations......................................................................85 

6.0 References.........................................................................................................................87 

List of Appendices 

Appendix A: No Action Alternative 

Appendix B: Exposure Calculation Worksheets 

Appendix C: Avian Species in Willapa Bay 

Appendix D: Terrestrial and Amphibious Wildlife in the Wallapa Bay Watershed 

Appendix E: Threatened and Endangered Species and Species of Concern within Washington 

State, and the Potential for Imazapyr Exposure from Spartina Treatment 

List of Tables 

Table 2-1: Spartina Distribution and Treatment in Washington State, 1997-2002. ..........................6 

Table 3-1: Product formulations of imazapyr. ..................................................................................23 

Table 3-2: Efficacy of Wipe-on Applications of Imazapyr and Glyphosate....................................25 

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Table 3-3: Examples of Aquatic Species Effectively Controlled by Imazapyr. ..............................26 

Table 3-4: Plants known to be resistant to the use of imazapyr........................................................26 

Table 3-5: Effect of rate, timing and spray volume in 2000 on the efficacy of imazapyr 

and glyphosate for smooth cordgrass control in Willapa Bay, WA. ........................30 

Table 3-6: Effects of herbicide rate and application spray volume in August 1999 on 

smooth cordgrass control in Willapa Bay, WA.........................................................31 

Table 3-7: Persistence and Bioconcentration of Imazapyr in Missouri and Florida Pond 

Water, Sediment and Resident Aquatic Biota...........................................................40 

Table 3-8: Hazard Classifications to Address Wildlife Risk from Herbicide Use...........................41 

Table 3-9: Acute and Subchronic Mammalian Toxicicity to Imazapyr...........................................42 

Table 3-10: Lethal and Sublethal Toxicity of Arsenal in Controlled Avian 

Experiments................................................................................................................44 

Table 3-11: Toxicity Classifications to Address Acute Risk to Aquatic Organisms from 

Chemical Use. ............................................................................................................47 

Table 3-12: 96-hour LC50 Values with 0.3 g juvenile rainbow trout exposed to 

imazapyr (Arsenal) or glyphosate (Rodeo) tank mixes. ...........................................48 

Table 3-13: Toxicity of Technical Grade Imazapyr and Arsenal* (with surfactant) to 

Algae and Aquatic Plants, as Established Through Controlled Product 

Registrant Studies.......................................................................................................51 

Table 3-14: Acetic Acid Toxicity to Ecological Receptors. .............................................................52 

Table 3-15: Effect of surfactant applied in September 1999 and 2000 on the efficacy of 

imazapyr for smooth cordgrass control in Willapa Bay, WA...................................53 

Table 3-16: Chemistry and Fate of Surfactants Potentially Used With Imazapyr and 

Glyphosate..................................................................................................................54 

Table 3-17: Toxicity of Surfactants With and Without Herbicide. ...................................................55 

Table 3-18: Acute toxicity of nonylphenol to aquatic biota. ............................................................56 

Table 4-1: Summary of Maximum and Average Imazapyr Detections in Relevant 

Environmental Media with Application Rates of 1.5 lbs a.i./acre............................61 

Table 4-2: Exposure Parameters for Addressing Risks from WSDA’s Imazapyr 

Applications for Spartina Control. ............................................................................63 

Table 4-3: Estimated Exposures to Terrestrial Wildlife Receptors from Imazapyr 

Applications (mg/kg-body wt)*.................................................................................69 

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Table 4-4: Estimated surfactant concentrations in estuary waters where Spartina could 

be treated with imazapyr, assuming constant surfactant rate of 1% in 

spray volume, complete solubility on incoming tide, and the “worst 

case” scenario of no adsorption on sediment or Spartina canopy. ...........................73 

Table 5-1: Hazard quotient calculations from estimated exposure doses of imazapyr to 

terrestrial wildlife.* ....................................................................................................75 

Table 5-2. Hazard Quotient Calculations from Estimated Aquatic Biota Exposures*....................80 

List of Figures 

Figure 2-1: Typical multi-clonal distribution of Spartina alterniflora in Willapa Bay..................5 

Figure 2-2: Typical Spartina alterniflora meadow in Willapa Bay. ...............................................5 

Figure 3-1: Chemical structures of the two forms of imazapyr. ....................................................24 

Figure 3-2: Box whisker graph of Spartina control as a function of rate of imazapyr and 

glyphosate use in experiments conducted in Willapa Bay, WA from 1997 to 2000 .28 

Figure 3-3: Box whisker graph of Spartina control as a function of dry time for imazapyr (1.68 

kg/ha) and glyphosate (8.4 kg/ha) use in experiments conducted in Willapa Bay,

WA form 1997 to 2000 ................................................................................................29 

Figure 3-4: Imazapyr Dissociation Under Different pH ................................................................32 

Figure 3-5: Degradation Pathway of Imazapyr ..............................................................................33 

Figure 3-6: Degradation Rate of Imazapyr in Sandy-Loam Soil ..................................................34 

Figure 3-7: Degradation of Imazapyr’s Initial Photodegradation Products CL 119060 and CL 

9140 in a controlled aerobic aquatic system using Missouri pond water .................36 

Figure 3-8: Residues of imazapyr in water and sediment from a Louisiana pond treated with

1.5 b a.i./acre ................................................................................................................37 

Figure 3-9: Persistence of imazapyr in estuarine waters of Willapa Bay following direct

application to an unvegetated tidal mud flat. Data represented are mean values of 

triplicate samples +/- SE ..............................................................................................38 

Figure 3-10: Persistence of imazapyr in estuarine sediment in Willapa Bay, WA after direct 

application to an unvegetated tidal mud flat. Data represented are mean values of 

triplicate samples +/- SE ..............................................................................................39 

Figure 3-11: Penetration distance in artificial beak tests .................................................................45 

Figure 3-12: Plasma sodium and gill ATPase activity of chinook salmon exposed to imazapyr . 50 

Figure 4-1: Mud-boat applicator used to apply herbicide in Willapa Bay to control Spartina....59 

Figure 4-2: Estimated water concentrations of imazapyr in tidal waters with no canopy 

interception and an application rate of 1.5 lbs/acre (0.68 kg/acre).............................60 

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Figure 4-3: Estimated plant residue concentrations over time, with an initial application rate of

1.5 lbs/acre. ...................................................................................................................65 

Figure 4-4: Estimated concentration of undiluted imazapyr in various spray volume applications.66 

Figure 5-1: Hazard quotient (HQ)estimation based on rainbow trout LC50 values of these 

surfactants as developed by Smith et al. (2003), where HQ = estimated water 

concentration/LC50. .....................................................................................................83 

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1.0 INTRODUCTION 

The Washington State Department of Agriculture (WSDA) has legislative mandate to control 

noxious and invasive weeds in the State of Washington. Noxious weeds are plants that when

established are highly destructive, competitive, or difficult to control by cultural or chemical 

practices (RCW 17.10.010). Cordgrass (Spartina spp.) a dominant invasive weed spreading 

throughout many of Washington’s most productive estuarine tide flats, is considered a noxious

weed under state code (WAC 16-750-011). Within the estuarine environments of Washington State 

only one herbicide, glyphosate (i.e. Rodeotm), is currently authorized for Spartina spp. control

(NPDS Permit # WAG-993000). Glyphosate is relatively non-toxic to animals and is effective on 

a wide range of plant species. However, its use to control cordgrass is hindered by drying times 

that limit its efficacy under the tidal conditions inherent to estuaries. It also requires higher

application rates than an alternative herbicide potentially available for use, imazapyr (i.e. 

Arsenaltm). The goals of this ecological risk assessment are therefore to 1) summarize current 

knowledge concerning the toxicity of of imazapyr to target and non-target organisms, 2) estimate

potential exposure to ecological receptors relevant to the aquatic (estuarine) environments where

the herbicide may be applied, and 3) characterize risks from that exposure to the individual species

and ecosystems where Spartina spp. is distributed. 

This ecological risk assessment should be considered supplemental to the original 1993

Environmental Impact Statement that evaluated the potential benefits and risks from the use of the

herbicide glyphosate (i.e. Rodeotm) and other mechanical management alternatives to control 

Spartina (WSDA 1993). Specifically, this assessment evaluates the risks to fish, wildlife and nontarget vegetation from the proposed use of imazapyr (i.e. Arsenaltm) to control smooth cordgrass in

estuarine waters of Washington State. Under the EPA Section 3 for Section 24(c) pesticide 

registration sought for imazapyr use in an estuary, applications would be made directly to the plant 

during low tide, and it is this scenario for which risks were considered. 

Appendix A to this supplement references the “no-action” alternative, which, in contrast to the 

earlier 1993 EIS, considers the “no-action” scenario to be a continuance of the current integrated

pest management scheme for Spartina control that involves the use of chemical (glyphosate) and 

mechanical control means. That is, “no action” in the current context and in the vernacular of

SEPA guidance, would constitute the environmental baseline upon which imazapyr use is

compared; the potential inclusion of imazapyr into WSDA’s integrated pest management scheme 

for Spartina control would therefore represent the “preferred alternative”. 

The outline and methods of the main body of this report reflect standard ecological risk assessment

guidelines (EPA 1996). Thus, the report begins with the “problem formulation”, which 

summarizes the scope of the problem, the need to consider alternative control mechanisms, and the 

approach to the assessment. The problem formulation is followed by the “hazard assessment”

which relates the current understanding of imazapyr’s environmental fate, and its toxicity to the 

range of target and non-target organisms where testing has been conducted. The hazard 

assessment is followed by the “exposure assessment,” where the pathways and doses possible for

imazapyr exposure to the representative biological receptors are evaluated. The exposure

assessment considers the threatened and endangered (T&E) avian and aquatic species where

imazapyr could be applied where data permit. If no toxicity data were available for species typical

of the estuarine environments where impazapyr use is proposed, then toxicity data from surrogate 

species were used. The use of surrogate species with similar dietary and/or behavior patterns has

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October 2003 Page 1 
been shown to provide a relatively reliable predictor of aquatic toxicity when toxicity data are 

lacking for species of greater relevance to specific areas (Sappington et al. 2000). 

Standard EPA and other test species were also used as surrogates to model potential exposure to 

terrestrial omnivores, herbivores and carnivores. It can be reasonably assumed that a similar

relationship as found with aquatic species sensitivity exists for other wildlife. However, only sitespecific risk assessments would be able to fully quantify risks to resident and migratory wildlife

receptors from chemical exposure. Notwithstanding, this assessment used surrogate species such

as the rat, rabbit, quail and mallard duck to gauge exposure to other wildlife that would be likely to

use estuarine habitat within Washington State. The rat provides a reasonable surrogate of an 

omnivore, the rabbit an exclusive herbivore, and the quail and duck provide surrogates of upland 

and wetland bird species, respectively. 

The effects of specific contaminants at the broader ecosystem level may also vary significantly

among ecosystems based on the physical and chemical properties of the chemicals themselves, and 

the unique combination of physical, chemical, and biological processes occurring in each

ecosystem. Such ecosystem differences can directly bring about differences in animal populations

or indirectly affect habitat. For example, wildlife populations resident to environments naturally

enriched with metals may tolerate a much higher concentration of metal exposure than naïve 

populations. That is, populations of exposed organisms may differ in their response to

contaminants depending on their natural tolerance to the chemical, their behavioral and life history 

characteristics (e.g., pre-exposure), the dose to which they are exposed, and the duration of

exposure. Furthermore, responses may be transient (and therefore reversible) or permanent

(irreversible). 

With the preceding discussion in mind, the objectives of this EIS supplement can be succinctly

summarized as follows: 

• To describe the toxicity hazards of imazapyr to marine and estuarine aquatic organisms, as 

known. 

• To describe the toxicity hazards of imazapyr to terrestrial and amphibious wildlife, as known. 

• To describe the toxicity hazards of imazapyr to non-target vegetation, as known. 

• To identify sensitive species that may be impacted in different regions where imazapyr could 

be applied. 

• To estimate (model) ecological receptor exposure (dose) by identifying complete and 

incomplete exposure pathways, taking into account environmental fate and transport through

both physical and biological means.

• To characterize the risk or threat to other environmental components potentially affected by 

imazapyr. 

• To compare risks from the potential use of imazapyr relative to the existing use of glyphosate

and other existing control methodologies, such that the existing control methods can be 

considered the “no action alternatives”. 

• To identify the method or integration of treatment methods, from review of new literature 

and WSDA’s existing program, that best controls Spartina spp. with the minimum amount of 

risk to the environment. 

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October 2003 Page 2 
2.0 PROBLEM FORMULATION 

To frame the scope of the Spartina problem in the State of Washington, the potential means to 

control it, and the probable risks and benefits of the use of Arsenaltm as a component in the

integrated management of the invasive weed, a thorough understanding of the scope of the problem 

is needed. This chapter defines the scope of the problem, the assessment endpoints that best

represent the management goals of the WSDA for Spartina control, the conceptual models we will 

use to consider exposure to relevant ecological receptors, and the methodology by which risks to

ecological receptors are quantified and characterized. 

Toxicology is the study of poisons. It examines and attempts to define the range in responses of an

organism or organisms to variable doses of a chemical or chemicals. Thus, the most important

factors regulating chemical toxicity are the exposure dose, the duration of exposure, and the

potency of the chemical. The genotype, and nutritional and physiological, state of an ecological

receptor at the time of exposure can also affect chemical toxicity. The introduction of chemicals

into an ecosystem can cause direct harm to organisms, or may indirectly affect their fitness—the 

ability of an animal to survive and produce viable offspring. The results of chemical exposure may

be immediately apparent or may become noticeable only after considerable delay. Recognizing the 

effects of exposure on animals may require analyses through a suite of measurement endpoints.

Measurement endpoints may include physiological, neurological, behavioral, endocrine-mediated, 

or a variety of other indicators that could be construed to play a role in the survival of the 

organism. 

Ecological risk assessment represents a branch of toxicology wherein the effects of putative

poisons are examined not only at the individual organismal level as outlined above, but also at the 

broader population and ecosystem level. Thus, the purpose of this ecological risk assessment is to

determine the nature, magnitude, and transience or permanence of observed or expected effects to 

animals and their habitat from exposure to imazapyr, based on WSDA’s projected application rates

and integrated pest management practices. The assessment relies heavily on ecological hazard 

studies that have been conducted over the past several years, product registration study results, and 

conservative deterministic exposure modeling at the organismal level. Effects at the organismal

level are presumed to be reflective of potential effects at the population level, though no 

quantitative measures of effect at the population level are calculated 

2.1 Overview of Spartina Infestation in Washington State 

Spartina is an invasive weed that inhabits tideflats, salt marshes and estuaries throughout

Washington State’s coastal areas. There are now four species of Spartina found in Washington’s 

waters, Spartina alterniflora, Spartina patens, Spartina anglica and a newly discovered species

found in the fall of 2001 in Grays Harbor and Island County, Spartina densiflora. Sometimes

referred to as cordgrass, Spartina alterniflora and Spartina patens are native to the Eastern and 

Gulf coasts of the United States and are integrated into these regions’ natural ecosystem processes.

Spartina anglica is an intentionally created hybrid. Spartina densiflora is native to South America

which has also invaded Humboldt Bay in California. In its native environments, the spread of

Spartina is controlled by natural biological agents, and by natural disturbance factors such as

hurricanes and tidal action. 

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October 2003 Page 3 
The potential of exotic species to change the physical structure and in so doing alter the ecological

functioning of the entire habitat unit has been well documented (Zipperer 1996). In several of 

Washington State’s coastal habitats, Spartina spp. outcompetes and displaces native vegetation and 

results in changes to local ecosystems by converting littoral mudflats to salt marshes. These 

changes threaten to impact native fisheries, shellfish beds, waterfowl migrations and other wildlife 

by reducing the habitats carrying capacity for these animals. 

2.1.1 Distribution

Spartina alterniflora typifies an invasive species by having a wide tolerance to habitat 

requirements, fast dispersal rate, clonal reproduction and few to no natural predators in its invaded

range (Zipperer 1996). Spartina alterniflora was most likely introduced to the Washington coast

when it was used in the packing of East Coast oysters for shipping during the late 1800s. In 

addition to the accidental introduction, S. alterniflora was also intentionally planted by a gun club

between 1941 and 1946 to stabilize bank erosion on their property in Padilla Bay. Spartina anglica

was also intentionally introduced, to stabilize dikes and provide forge for cattle in Port Susan Bay. 

The pathways of introduction for both Spartina patens and the newly discovered S. densiflora are

not known at this time. 

Spartina species have spread throughout Washington State’s coastal counties. Infestations range

from only a few square feet to more than 6,800 solid acres. Currently there are an estimated 7,500

solid acres interspersed amongst 20,000 total invaded acres in Washington’s coastal habitat. Table 

2-1 summarizes the locations, size and recent treatments of Spartina sp. found in Washington State

(WSDA Legislative Report 2002). Figures 2-1 and 2-2 demonstrate typical clone and field 

colonies (meadows) of Spartina, as occurring today in Willapa Bay. 

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October 2003 Page 4 
Figure 2-1: Typical multi-clonal distribution of Spartina alterniflora in Willapa Bay.

(Source: WSDA 2003.)

Figure 2-2: Typical Spartina alterniflora meadow in Willapa Bay.

(photo credited to K. Patten, by permission.)

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October 2003 Page 5 
Table 2-1: Spartina Distribution and Treatment in Washington State, 1997-2002.

County Spartina Present in 2002 Spartina Treated, 1997 - 2002 2002 Treatment Methods 

Pacific (Willapa Bay) Over 6,800 solid acres 

spread over > 15,000

acres 

1997 –approx. 742 solid acres 

1998 –approx. 450 solid acres 

1999 –approx. 600 solid acres 

2000 –approx. 800 solid acres 

2001 –approx. 900 solid acres 

2002 –approx. 1804 solid acres 

Mow/herbicide, herbicide, seedling 

removal, various mechanical control. 

Grays Harbor Scattered clones and

seedlings 0.25 acres each

1997 through 2002 - all treated Herbicide, seedling removal, mow

Snohomish Approx. 350 solid acres 

spread over >4,500 acres 

1997 –approx. 89 solid acres 

1998 –approx. 126 solid acres 

1999 –approx. 90 solid acres 

2000 –approx. 158 solid acres 

2001 –approx. 75 solid acres 

2002 –approx. 238 solid acres 

Mow/herbicide, herbicide, seedling 

removal, dig, mechanically crush, mow. 

Island Approx. 350 solid acres 

spread over >1,000 acres 

1997 –approx. 250 solid acres 

1998 –approx. 160 solid acres 

1999 –approx. 155 solid acres 

2000 –approx. 130 solid acres 

2001 –approx. 72 solid acres 

2002 –approx. 180 solid acres 

Mow/herbicide, herbicide, seedling 

removal, mechanically crush, mow.

Skagit Approx. 40 solid acres 

spread over >2,000 acres 

1997 –approx. 91 solid acres 

1998 –approx. 57 solid acres 

1999 - all treated

2000 –approx. 60 solid acres 

2001 –approx. 33 solid acres 

2002 –approx. 37 solid acres 

Mow/herbicide, herbicide, seedling 

removal, dig, mow. 

Clallam 1 infestation < 0.001 acres 

in size 

1997 – treated twice 

1998 – treated three times 

1999 – treated twice 

2000 – treated three times 

2001-02 – treated four times 

Dig 

Jefferson 14 infestations - approx. 

0.01 solid acres total 

1997 - all treated

1998-2000- all treated twice

2001-02 – all treated three times 

Mow, mow/herbicide, dig, seedling 

removal

Kitsap 8 infestations - approx. 1 

solid acre total 

1997 - all but 2 tribal sites 

1998 - all treated once 

1999 - all treated twice 

2000-01 - all treated once 

2002 - all treated twice 

Mow, mow/herbicide, dig, seedling 

removal

King 2 infestations - single

clones and a few seedling 

1997 – monitored

1998-99 - all treated once 

2000-02 - all treated twice 

Dig 

San Juan Re-growth found at one 

site. 2 other sites clean for

four consecutive years 

97 - all treated 

98 - all treated 

99 – monitored 

2000-02 - all treated once 

Survey, dig 

From Washington State Department of Agriculture’s Report to the Legislature, Progress of the 2002 Spartina Eradication Program, 

December 15, 2002 

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2.1.2 Biology 

Spartina is a rhizomatous perennial grass that can proliferate from either sexual reproduction or

vegetative propagation. Shoots sprout from below ground rhizomes in the spring and reach a height 

of three to six feet by mid-summer. Spartina flowers from late June to October; however, not all

populations within Washington State flower. Sexual reproduction may require a sufficient 

underground biomass to trigger. Populations that do not flower depend on vegetative propagation 

and/or lateral growth to spread. Temperature, photoperiod soil temperature, and soil salinity have

been shown to influence occurrence and timing for the populations that do flower. Salinity has also 

been shown to affect growth rate, seedling development and spatial zonation (Feist 1999). 

Dispersal of Spartina is accomplished when water currents, animals or humans transport seed, 

rhizome pieces or entire plants to new locations. Humans are the most prevalent cause of Spartina

dispersal, either through intentionally plantings or accidental spread. Waterfowl and other birds are 

known to ingest seeds and rhizomes, spreading material through their feces. Water spread seeds

and rhizomes fragments via natural currents and tidal actions. These mechanisms of dispersal

create great difficulties in controlling the spread of the weed. 

When new plants are established through either seeds or vegetative propagation, survival appears 

to be linked to competition with other plants. Low light levels caused from other plants may inhibit 

survival. Seed and vegetative propagation seems to be important in the colonization of disturbed or

bare areas. In contrast, growth of established colonies out- competes native plant species resulting 

in a nearly complete monoculture of Spartina plants in invaded habitats. These colonies or clones

reproduce by the lateral spread of underground rhizomes and aboveground tillers. 

Spartina alterniflora, S. patens, and S. anglica become dormant during the winter and die back. 

However, Spartina densiflora does not become dormant and produces new plant material

throughout the year. In addition, the seed viability of S. densiflora appears to be much higher than

the other species of Spartina. These traits may potentially allow S. densiflora to invade new areas 

more rapidly than the other Spartina species in Washington State. 

In its native habitats S. alterniflora functions as essential feeding grounds and nursery areas for 

numerous species of invertebrates, fish, shorebirds, migratory waterfowl, and small mammals. In 

contrast S. alterniflora alteration of mud flats could eliminate critical foraging habitat of juvenile 

salmonids, flatfish, shorebirds, and migratory waterfowl. In addition, composition and abundance

of benthic invertebrates may be substantially altered by cordgrass colonization of mudflats because

these species are strongly influenced by the physical environment (Zipperer 1996). 

2.2 Ecological Receptors, Community Descriptions, and Threatened and 

Endangered Species In Primary Areas Where Spartina is Distributed 

As demonstrated in Table 2-1, Spartina is found in multiple locations within Puget Sound, and on

the Pacific Coast within Willapa Bay. Most areas of Spartina distribution in Puget Sound are

localized and can be controlled by mechanical means (see Table 2-1). The two areas of varying 

size and distribution, Willapa Bay and Paddila Bay, require the use of chemical control means to

achieve the goal of eradication, and chemical control must remain a viable alternative for all areas 

where Spartina is found if mechanical means prove ineffective in the future. With this 

understanding, an overview of the ecological communities and ecological receptors in Willapa and

Padilla bays is provided below. The Willapa Bay and Padilla Bay ecosystems share many 

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similarities with the other areas Spartina has colonized in Washington State, although due to their

size, the ecosystems support a greater diversity of species. These habitats, as well as other coastal

habitats of Washington where smaller colonies of Spartina have established, support several 

priority species, some of which are listed as threatened or endangered (T&E) under the Endangered 

Species Act (ESA). Thus, this section also provides a summation of the T&E species in

Washington State, and the potential for these species to occur in areas where imazapyr treatments 

of Spartina could occur. 

Willapa Bay

Willapa Bay is located in the southwestern corner of the Washington coast. It is approximately 38

km long and 8 km wide (Gringas et al. 2000). At high tide, the aquatic environment of Willapa 

Bay is approximately 88,000 acres, however almost half of the Bay is drained at low tide (Cohen et 

al. 2001). Willapa Bay in almost fully enclosed by the Long Beach Peninsula, a 30 km-long 

barrier spit that was formed by the deposition of Columbia River sediments. The Willapa Basin 

has received an average of 85 inches of rain per year over the past 75 years of record

(http://www.tidepool.org/wiscweb/wisc98nw2.html). Willapa Bay has a drainage basin of

approximately 2,550 square miles, and the main tributaries that drain into the bay include the 

North, Willapa, and Naselle rivers. The Palix River is on of the minor contributors to the mean 

daily runoff. Mean daily runoff to Willapa Bay represents approximately < 0.05 percent of the bays

total volume. 

Willapa Bay is Washington’s largest outer coast estuary (Cohen et al. 2001). Willapa Bay is a 

largely unaltered environment, however it has been significantly impacted by the colonization of 

non-indigenous/exotic species. Of the 892 vascular plants in the Willapa Basin (which includes 

headwater habitat outside of the brackish estuary) approximately 250 species have been 

introduced. Similarly, 30 of the 473 species of vertebrates identified in the basin have been

introduced (www.tidepool.org/wiscweb/wisc98nw2.html). Approximately 34 exotic aquatic plant 

and animal species were recently identified within the Willapa Bay estuary during a 2000 research 

expedition sponsored by the Washington State Department of Natural Resources (WDNR)

Nearshore Habitat Program (Cohen et al. 2001). Within the estuary habitat of the basin, the two 

most significant plant species introduced are the Japanese eelgrass (Zostera japonica) and the

smooth cordgrass (Spartina alterniflora). 

The most problematic exotic species found within Willipa Bay is the smooth cordgrass. This 

invasive species is outcompeting native aquatic vegetation for space and nutrient resources.

Spartina also degrades habitat for aquatic, terrestrial and aviananimal species utilizing Willapa 

Bay. Between 1994 and 1997, Spartina populations expanded at a rate of 485 percent within the

southern portion of Willapa Bay (Willapa Bay Estuary 2001). 

In addition to the major issues created by the introduction of Spartina, numerous aquatic

invertebrate animal species have been introduced intentionally or inadvertently into Willapa Bay 

during the past century. The degree to which these introductions have displaced native species is

less understood than the displacement caused by Spartina. Some of the known non-native

introductions are tabulated in Table 2-2. 

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Table 2-2: Common Non-native Invertebrates in Willapa Bay.

General Taxon Species 

First Puget 

Sound 

Record

Hobsonia florida 1940 

Polydora cornuta 1932 

Pseudopolydora bassarginensis 2000 

Pseudopolydora kempi japonica 1951 

Annelida (polychaeta) 

Streblospio benedicti 1932 

Crepidula fornicata 1905 

Illyanassa obsoleta 1907 

Ocinebrellus inornatus 1924 Molluska (prosobranchia) 

Urosalpinx inornatus 1890 

Crassostrea gigas 1875 

Mya arenaria 1874 

Neotrapezium liratum 1924 

Petricolaria pholadiformis 1927 

Molluska (bivalve) 

Venerupis philippinarum 1924 

Eusariella zostericola 1953 

Balanus improvisus 1853 

Nippoleucon hinumensis 1979 

Limnoria tripunctata 1871 or 1875 

Ampithoe valida 1941 

Corophium acherusicum 1905 

Corophium insidiosum 1915 

Grandidierella japonica 1966 

Jassa marmorata 1938 

Anthropoda (crustacea) 

Melita nitida 1938 

Entoprocta (bryozoa) Bowerbanki gracilis 1923 

Botrylloides violaceus 1973 

Urochordata (ascidiacea) Botryllus schlosseri 1944-47 

Molgula manhattensis 1949 

Porifera Clathria prolifera 1945-49 

Cnidaria (hydrozoa) Cordylophora caspia 1920 

Cniidaria (anthozoa) Diadumene lineata 1906 

Source: Cohen et al. 2001 

Several of the resident shellfish species in Willapa Bay support substantial commercial harvest 

and/or farming industries (Table 2-3). The most significant species include the Pacific Oyster 

(Crassostrea gigas) and the native Dungeness crab (PNCERS 1998). 

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Table 2-3: Commercial Shellfish Species within Willapa Bay.

Common Name Scientific Name 

Type of

Species 

Pacific oyster Crassostrea gigas Oyster

Dungeness crab Cancer magister Crab 

Red rock crab Plagusia chabrus Crab 

Geoduck Panopea abrupta Clam 

Quahog (hardshell) Arctica islandica Clam 

Softshell clam Mya arenaria Clam 

Native littleneck Protothaca staminea Clam 

Cherrystone Mercenaria mercenaria Clam 

Source: PNCERS 1998 

Anadromous salmonids use Willapa Bay’s major tributaries for migration, spawning, incubation 

and early-rearing (Table 2-4). Habitat within Willapa Bay is also important habitat for larval and 

juvenile marine and anadromous fish rearing. It is, “arguably the most important nursery estuary 

on the coast for juvenile English sole” (B. Dumbauld, WDFW, personal communication to Wendy 

Sue Wheeler, WSDA, 11/14/2000). Pacific herring spawn on the eelgrass beds in the early spring, 

and the estuary also supports lesser-known smelt and sand lance runs; all three of these species are

important forage fish for Pacific salmon. Anchovy, salmon and sturgeon have supported

commercial fisheries in the outer bay in the past. 

Willapa Bay is also a major migration stopover location for shorebirds in the spring and winter 

(Willapa National Wildlife 2001). An estimated 100,000 to 1,000,000 shorebirds stop to feed in the 

mudflats of Willapa Bay and other coastal regions of Washington State during the spring. 

However, spring and winter peak shorebird numbers have been declining by 54 and 67 percent,

respectively since 1991 due to infestation of Spartina. 

The distribution of ducks within Willapa Bay was modeled by Willapa National Wildlife (2001).

The hierarchy of distribution within Willapa Bay according to mid-winter aerial waterfowl surveys

is: South Bay (47.1%) > East Bay (28.6%) > North Bay (18.8%) > West Bay (4.2%) > Peninsula

(1.2%). The most significant region for ducks, the South Bay, is also harbors the greatest density 

of Spartina. A summation of some of the common avian species found within Willapa Bay is 

provided in Appendix C. (USFWS 1991). 

Willapa Bay, its surrounding wildlife refuge, and the extensive contigous lowland forests also

support a diverse assemblage of terrestrial and amphibious wildlife. Some 53 species of mammals 

and 19 herptiles (reptiles and amphibians) have been reported, as summarized in Appendix D 

(USFWS 1991). 

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Table 2-4: Anadromous Salmonid Distribution and Utilization within Willapa Bay 

Tributaries.

River Species Run Primary Use River Miles Used % of Stream 

Used 

Migration 0.0-7.3 20% Chinook salmon Fall

Spawning/ Rearing 7.3-31.2 63%

Migration 0.0-7.2 19% Coho salmon N/A 

Spawning/ Rearing 7.2-26.8 52%

Migration 0.0-7.4 20% Steelhead Winter

Spawning/ Rearing 7.4-30.2 61%

Migration 0.0-9.9 26%

Rearing/ Migration 9.9-15.2 14%

Spawning/ Rearing 15.2-25.2 27%

Naselle River

Chum salmon N/A 

Migration 25.2-25.6 1%

Migration 0.0-0.3 1% Chinook salmon Fall

Spawning/ Rearing 0.3-59.3 98%

Migration 0.0-0.3 1% Coho salmon N/A 

Spawning/ Rearing 0.3-60.0 99%

Migration 0.0-0.6 1%

Steelhead Winter Rearing/ Migration 0.6-22.0 36%

Spawning/ Rearing 22.0-60.0 63%

North River 

Chum salmon N/A Migration 0.0-5.8 10%

Chinook salmon Fall Migration 0.8-4.0 34%

Coho salmon N/A Migration 0.8-4.0 34%

Steelhead Winter Migration 0.8-4.0 34%

Palix River 

Chum salmon N/A Migration 0.8-4.0 34%

Migration 0.0-7.5 16% Chinook salmon Fall

Spawning/ Rearing 7.5-41.3 72%

Migration 0.0-5.5 12%

Rearing/ Migration 5.5-5.8 1%

Spawning/ Rearing 5.8-41.8 76%

Coho salmon N/A 

Migration 41.8-44.1 5%

Migration 0.0-5.5 12%

Steelhead Winter Rearing/ Migration 5.5-28.2 48%

Spawning/ Rearing 28.2-41.3 28%

Migration 0.0-28.3 60%

Spawning/ Rearing 28.3-31.8 8%

Willapa River 

Chum salmon N/A 

Migration 31.8-36.0 9%

Willapa Bay Green sturgeon N/A Spawning/ Rearing N/A N/A 

Source: StreamNet 2003 

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Padilla Bay

Padilla Bay is an 11,000 acre shallow-water bay in north Puget Sound, incorporating the deltas of 

the Skagit and Samish Rivers. Padilla Bay is delineated by the saltwater edge of the North Fork

Skagit River delta in Puget Sound. Padilla Bay is approximately 8 miles long and 3 miles wide 

(Padilla Bay NERR 2002). The Skagit River provides the majority of the freshwater and sediment

resources to the bay. The bottom of Padilla Bay is very shallow due to sediment transport from the 

Skagit River, which creates a broad tidal flat during low tide and flooded during high tide (Padilla

Bay NERR 2002). 

Eelgrass meadows occupy nearly 8,000 acres of the bay, and are made up primarily of two species: 

native eelgrass (Zostera marina) and a non-native species (Zostera japonica). The eelgrass

meadows stabilize the mud-flat substrate that dominates the bay, and provide food and shelter for

various fish and wildlife. Eelgrass and algae are the main primary producers within Padilla Bay

(Thom 1988). The salt marsh associated with Padilla Bay was diked and drained before 1900 for 

farm land, leaving a small fringe of the salt marsh that includes species such as salt grass 

(Distichlis spicata), salt brush (Ariplex patula), pickleweed (Salicornia virginica), and seaside 

arrowgrass (Triglochin maritimum). 

The salt marsh and mudflat of Padilla Bay has problems with two non-native invasive species of 

cordgrass (Spartina alterniflora and Spartina anglica) that are competing for resources held by 

native species (WSDA 2000). S. alterniflora (smooth cordgrass) was first introduced into Padilla 

Bay in the early 1940s and by 1979 approximately 3.5 acres were reported in Padilla Bay; S. 

alterniflora was reported to have increased to 17 acres by 1997 (WSDA 2000). S. anglica

(common cordgrass) formed from allopolyploidy of the sterile hybrid S. X townsendii in England. 

S. anglica has strong hybrid vigor and has taken over approximately 25,000 acres of intertidal salt 

marsh on the British coast within the past 100 years (WSDA 2000). S. anglica was reported in the 

Puget Sound by 1979 in an estimated coverage of 15 acres; by September 1997, approximately

1,000 solid acres was reported within North Puget Sound (over 8,000 acres of this region was

impacted). 

Aquatic organisms found within Padilla Bay include crabs, shrimp, mud snail, and various

organisms that are supported in salt marsh/mud flat habitat. Extensive eelgrass meadows in the

bay provide excellent habitat for finfish such as salmon, perch, and herring, but also many 

invertebrate species (e.g. worms, shrimp, clams). These species in turn support great blue heron,

eagle, otter, and seal populations. The eelgrass meadows of the Padilla Bay estuary provide 

suitable habitat for many different life stages of aquatic organisms. For example, young 

Dungeness crabs, one of the most economically important aquatic organisms in Padilla Bay, utilize 

intertidal cobble found within the eelgrass meadows (Dinnel et al. 1986). 

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Table 2-4: Marine Invertebrate Species within Padilla Bay. 

Phylum Common Name Scientific Name Exotic

Nemertea Sand nemertean Cerebratulus californiensis no 

Nemertea Green nemertean Emplectonema gracile no 

Nemertea Restless worm Paranemertes peregrina no 

Annelida Lugworm Abarenicola pacifica no 

Annelida Rough-skinned lugworm Abarenicola claparedii no 

Annelida Thread worm Notomastus tenuis no 

Cnidaria Sea pen Abietinaria sp. no 

Cnidaria Orange-striped jellyfish Gonionemus vertens no 

Cnidaria Aggregate anemone Anthopleura elegantissima no 

Cnidaria Brooding anemone Epiactis prolifera no 

Cnidaria Tealia Tealia sp. no 

Cnidaria Stalked jellyfish Haliclystus auricula no 

Ctenophora Sea gooseberry Pleurobrachia bachei no 

Brachiopoda Lamp shell Terebratalia transversa no 

Echinodermata Blood star Henricia leviuscula no 

Echinodermata Six-rayed sea star Leptasterias hexactis no 

Echinodermata Purple star Pisaster ochraceus no 

Echinodermata Sunflower star Pycnopodia helianthoides no 

Echinodermata Green sea urchin Strongylocentrotus droebachiensis no 

Echinodermata Red sea cucumber Cucumaria miniata no 

Echinodermata White sea cucumber Eupentacta quinquesemita no 

Chaetognatha Arrow worm Sagitta elegans no 

Chordata Hairy sea squirt Boltenia villosa no 

Chordata Broad base sea squirt Cnemidocarpa finmarkiensis no 

Chordata Warty sea squirt Pyura haustor no 

Mollusca Mossy chiton Mopalia muscosa no 

Mollusca Large variegated limpet Notoacmea persona no 

Mollusca Plate limpet Notoacmea scutum no 

Mollusca Finger limpet Collisella digitalis no 

Mollusca Shield limpet Collisella pelta no 

Mollusca Limpet Unidentified sp. no 

Mollusca Spindle whelk Searlesia dira no 

Mollusca Chinese hat Calyptraea fastigiata no 

Mollusca Hooked slipper shell Crepidula adunca no 

Mollusca Slipper shell Crepidula sp. no 

Mollusca Screw snail Bittium sp. no 

Mollusca Hairy shell Trichotropis sp. no 

Mollusca Amphissa Amphissa sp. no 

Mollusca Keyhole limpet Diodora aspera no 

Mollusca Chink shell Lacuna variegata no 

Mollusca Sitka periwinkle Littorina sitkana no 

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October 2003 Page 13
Phylum Common Name Scientific Name Exotic

Mollusca Checkered periwinkle Littorina scutulata no 

Mollusca Cowry? Cypraeolina pyriformis no 

Mollusca Japanese hornmouth Ocenebra inornata (=japonica) yes 

Mollusca Atlantic oyster drill Urosalpinx cinerea yes 

Mollusca Basket shell Nassarius fraterculus yes 

Mollusca Lean basket shell Nassarius mendicus no 

Mollusca Lewis' moon snail Polinices lewisii no 

Mollusca Turret shell Batillaria attramentaria yes 

Mollusca Japanese false cerith Batillaria zonalis yes 

Mollusca Wrinkled thais Thais lamellosa no 

Mollusca Blue top shell Calliostoma ligatum no 

Mollusca Puppet margarite Margarites pupillus no 

Mollusca Taylor's sea slug Phyllaplysia taylori no 

Mollusca Bubble shell Acteocina sp. no 

Mollusca Barrel bubble Acteocina harpa (Retusa harpa) no 

Mollusca Bubble shell Haminoea sp. no 

Mollusca Blister paper bubble Haminoea vesicula no 

Mollusca Barrel bubble Cylichna sp. no 

Mollusca Odostome Odostomia sp. no 

Mollusca Opalescent nudibranch Hermissenda crassicornis no 

Mollusca Sculptured nut clam Acila castraensis no 

Mollusca Heart cockle Clinocardium nuttallii no 

Mollusca Japanese oyster Crassostrea gigas yes 

Mollusca Dipper clam Lyonsia striata no 

Mollusca Polluted macoma Macoma inquinata no 

Mollusca Bent-nosed clam Macoma nasuta no 

Mollusca Sand clam Macoma secta no 

Mollusca Eastern soft-shell clam Mya arenaria yes 

Mollusca Blunt soft-shell clam Mya truncata no 

Mollusca Blue mussel Mytilus edulis no 

Mollusca purple varnish clam Nuttallia obscurata yes 

Mollusca Rock oyster Pododesmus macroschisma no 

Mollusca Native littleneck clam Protothaca staminea no 

Mollusca Washington clam Saxidomus giganteus no 

Mollusca Butter clam Saxidomus nuttalli no 

Mollusca Horse clam Schizothaerus nuttallii no 

Mollusca Jackknife clam Solen sicarius no 

Mollusca Japanese littleneck clam Venerupis philliparum yes 

Mollusca White tellen Tellina modesta no 

Mollusca Horse clam Tresus capax no 

Mollusca Dentalium Dentalium rectuis no 

Arthropoda Horse barnacle Balanus cariosus no 

Arthropoda Smooth acorn barnacle Balanus crenatus no 

Arthropoda Acorn barnacle Balanus glandula no 

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October 2003 Page 14
Phylum Common Name Scientific Name Exotic

Arthropoda Eelgrass isopod Idotea resecata no 

Arthropoda Olive-green isopod Idotea wosnesenskii no 

Arthropoda Oregon pill bug Gnorimosphaeroma oregonense no 

Arthropoda Beach hopper Orchestia traskiana no 

Arthropoda Skeleton shrimp Caprella laeviscula no 

Arthropoda Coon-striped shrimp Pandalus danae no 

Arthropoda Gray shrimp Crangon nigricauda no 

Arthropoda Short-spined shrimp Heptacarpus brevirostrus no 

Arthropoda Ghost shrimp Callianassa californiensis no 

Arthropoda Mud shrimp Upogebia pugettensis no 

Arthropoda Porcelain crab Petrolisthes eriomerus no 

Arthropoda Hermit crab Pagurus granosimanus no 

Arthropoda Hairy hermit crab Pagurus hirsutiusculus no 

Arthropoda Decorator crab Oregonia gracilis no 

Arthropoda Spider crab Pugettia gracilis no 

Arthropoda Kelp crab Pugettia producta no 

Arthropoda Dungeness crab Cancer magister no 

Arthropoda Red rock crab Cancer productus no 

Arthropoda Graceful cancer Cancer gracilis no 

Arthropoda Purple shore crab Hemigrapsus nudus no 

Arthropoda Green shore crab Hemigrapsus oregonensis no 

Arthropoda Pea crab Pinnixa occidentalis no 

Arthropoda Pea crab Pinnixa schmitti no 

Arthropoda Burrow crab Pinnixa tubicola no 

Arthropoda Helmet crab Telmessus cheiragonus no 

Arthropoda Sea spider Halosoma viridintestinale no 

Source: S. Riggs, Personal Communication, Padilla Bay National Estuarine Research Reserve, 2003

The complex of aquatic and intertidal habitats supported in Padilla Bay and its conjoined 

freshwater deltaic environments support the early rearing and the saltwater/freshwater

physiological transitions of substantial anadromous salmonid stocks (Table 2-5). In addition, over

50 other resident fish species have been reported in the bay (Table 2-5). Both herring and smelt

use the eelgrass meadows of Padilla Bay for spawning, and both species are significant salmonid

forage species. 

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Table 2-5. Anadromous and Resident Fish Species Found within Padilla Bay and 

Associated Tributaries.

Common name Scientific Name 

Anadromous/ 

Resident 

Pink salmon Oncorhynchus gorbuscha Anadromous 

Chum salmon Oncorhynchus keta Anadromous 

Coho salmon Oncorhynchus kisutch Anadromous 

Sockeye salmon Oncorhynchus nerka Anadromous 

Chinook salmon Oncorhynchus tshawytscha Anadromous 

Coastal cutthroat trout Salmo clarki clarki Anadromous 

Steelhead Oncorhynchus mykiss Anadromous 

Dolly varden/ bull trout Salvelinus malma Anadromous 

Spiny dogfish Squalus acanthias Resident 

Big skate Raja binoculata Resident 

Ratfish Hydrolagus colliei Resident 

Pacific herring Clupea harengus pallasi Resident 

Northern anchovy Engraulis mordax mordax Resident 

Surf smelt Hypomesus pretiosus pretiosus Resident 

Longfin smelt Spirinchus thaleichthys Resident 

Northern lampfish Stenobrachius leucopsarus Resident 

Plainfin midshipman Porichthys notatus Resident 

Northern clingfish Gobiesox maeandricus Resident 

Pacific tomcod Microgadus proximus Resident 

Red brotula Brosmophycis marginata Resident 

Blackbelly eelpout Lycodopsis pacifica Resident 

Tube-snout Aulorhynchus flavidus Resident 

Three-spined stickleback Gasterosteus aculeatus Resident 

Bay pipefish Syngnathus griseolineatus Resident 

Shiner perch Cymatogaster aggregata Resident 

Pile perch Rhacochilus vacca Resident 

Striped seaperch Embiotoca lateralis Resident 

Pacific sandfish Trichodon trichodon Resident 

Northern ronquil Ronquilus jordani Resident 

(Pacific) snake prickleback Lumpenus sagitta Resident 

Bluebarred prickleback Plectobranchus evides Resident 

Black prickleback Xiphister atropurpureus Resident 

Penpoint gunnel Apodicthys flavidus Resident 

Crescent gunnel Pholis laeta Resident 

Saddleback gunnel Pholis ornata Resident 

Pacific sand lance Ammodytes hexapterus Resident 

Yellowtail rockfish Sebastes flavidus Resident 

Sharpchin rockfish Sebastes zacentrus Resident 

Rock greenling Hexagrammos lagocephalus Resident 

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October 2003 Page 16
Common name Scientific Name 

Anadromous/ 

Resident 

Whitespotted greenling Hexagrammos stelleri Resident 

Lingcod Ophiodon elongatus Resident 

Padded sculpin Artedius fenestralis Resident 

Silverspotted sculpin Blepsias cirrhosus Resident 

Sharpnose sculpin Clinocottus acuticeps Resident 

Buffalo sculpin Enophrys bison Resident 

Soft sculpin Gilbertidia sigalutes Resident 

Pacific staghorn sculpin Leptocottus armatus Resident 

Great sculpin Myoxocephalus polyacanthocephalus Resident

Sailfin sculpin Nautichthys oculofasciatus Resident 

Tadpole sculpin Pyschrolutes paradoxus Resident 

Grunt sculpin Rhamphocottus richardsoni Resident 

Ribbed sculpin Triglops pingeli Resident 

Cabezon Scorpaenichtyhys marmoratus Resident 

Sturgeon poacher Agonus acipenserinus Resident 

Tubenose poacher Pallasina barbata aix Resident 

Smooth alligator fish Anoplagonus inermis Resident 

Pacific spiny lumpsucker Eumicrotremus orbis Resident 

Spotted snailfish Liparis callyodon Resident 

Ribbon snailfish Liparis cyclopus Resident 

Marbled snailfish Liparis dennyi Resident 

Tidepool snailfish Liparis florae Resident 

Showy snailfish Liparis pulchellus Resident 

Speckled sanddab Citharichthys stigmaeus Resident 

Arrowtooth flounder Atheresthes stomias Resident 

Starry flounder Platichthys stellatus Resident 

Butter sole Isopsetta isolepis Resident 

Rock sole Lepidopsetta bilineata Resident 

Slender sole Lyopsetta exilis Resident 

Dover sole Microstomus pacificus Resident 

English sole Parophrys vetulus Resident 

Sand sole Psettichthys melanostictus Resident 

Source: S. Riggs, Personal Communication, Padilla Bay National Estuarine Research Reserve, 2003

Padilla Bay is a reserve for migratory avian species in the winter (approximately 50,000 ducks, 

covering 26 species), as well as resident species (Padilla Bay NERR 2002). Resident species

include great blue heron, dunlin (a shorebird), bald eagle, peregrine falcon, merlin, and snowy owl.

There are approximately 240 species of birds that utilize Padilla Bay as either a foraging resource, 

nesting area, or migratory route. 

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Terrestrial mammals in the Padilla Bay reserve include black-tailed deer, raccoon, skunks, coyote,

muskrat, and long-tailed weasel; marine mammals that use Padilla Bay include harbor seals, and

occasionally California sea lions and porpoises (Table 2-6). 

Table 2-6: Terrestrial and Aquatic Mammalian Species in the Padilla Bay Region. 

Order Common name Scientific name Exotic

Marsupiala Virginia opossum Didelphis virginiana yes 

Vagrant shrew Sorex vagrans no Insectivora 

Mole Unidentified sp. no 

Chiroptera Bat Myotis spp. no 

Lagomorpha Eastern cottontail Sylvilagus floridanus yes 

Douglas' squirrel Tamiasciurus douglasii (i or ii) no 

Northern flying squirrel Glaucomys sabrinus no 

Beaver Castor canadensis no 

Townsend's vole Microtus townsendii (i or ii) no 

Muskrat Ondatra zibethicus (zibethica?) no 

Rodentia 

Deer (white-footed) mouse Peromyscus maniculatus no 

Coyote Canis latrans no 

Red fox Vulpes vulpes (fulva?) no 

Raccoon Procyon lotor no 

Striped skunk Mephitis mephitis no 

River otter Lutra candensis no 

Carnivora 

Longtailed weasel Mustela frenata no 

Artiodactyla Mule (black-tailed) deer Odocoileus hemionus columbianus no 

Cetacea Porpoise Unidentified sp. no 

Harbor seal Phoca vitulina no Pinnipedia 

Sea lion Unidentified sp. no 

Source: S. Riggs, Personal Communication, Padilla Bay National Estuarine Research Reserve, 2003

Specific Receptors Examined for Exposure 

Plants typical to the environments where imazapyr could be used to control Spartina include such

species as eelgrass (Zostera marina and Zostera japonica), and a variety of algal species such as

sea lettuce (Ulva sp.). Animals could include ungulates such as deer, elk and rabbit, omnivores

such as raccoons, terrestrial carnivores such as bobcat and coyote, avian species such as osprey, 

eagles and gulls; reptiles such as turtles; amphibians such as frogs; and insects such as mosquitoes. 

Obligate aquatic animal species include the array of Pacific salmonids native to Washington’s

waters, such as coho salmon, but also other fish species such as juvenile flatfish (Pleuronectidae), 

juvenile sturgeon (Ascipenseridae), and bullhead (Cottidae). Additional aquatic species potentially 

exposed to imazapyr include the vast list of benthic and mobile invertebrates common to the 

intertidal zone of Washington’s estuaries such as dungeness and rock crab (Cancer spp). As 

opposed to modeling exposure to all of these (and other) possible plants and animals that could be

potentially exposed to imazapyr from Spartina treatment, we evaluated exposure in select

surrogate “guilds”. 

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Receptor “guilds,” species with similar life histories or niches in the environment, are used to 

estimate exposure rather than estimating exposure for each individual species where a chemical 

could be applied. The assumption of this approach is that the general characteristics of each guild

will provide risk estimates that are representative of the entire guild. As such, each guild can be

extrapolated more broadly than single species estimates. The underlying concept is that each 

receptor falls into a group of potential receptors that function in similar ecological niches or

"guilds." For example, many species of heron and egret feed on small fish and invertebrates and 

require trees for roosts. As such, herons and egrets display similar life histories and would be

anticipated to have similar exposures to imazapyr. A single surrogate, such as the great blue heron, 

for which reliable life-history information is available, may be used for calculating risk and the

results may then be extrapolated to the guild as a whole. This approach allows the risk assessment

to directly evaluate species for which the best exposure information is available. This approach

also allows results to be extrapolated to a broader range of potential receptors, thereby maximizing 

data usage and applicability of results. 

Surrogate species were selected in each identified receptor guild. The selected surrogates have

been studied sufficiently to enable risk calculations to be made even though a surrogate itself may 

not necessarily be present within the study area (e.g., mallard). All of the other receptors are

present in Washington State and are representative of feeding guilds present. The fundamental 

assumption that was made in this study was that if negligible risk is determined for the surrogate 

species, then the entire guild is protected. 

Specific wildlife receptor guilds were selected based on the evaluation of exposure pathways and 

the possibility that a given receptor could come into contact with imazapyr applied for Spartina 

control. The receptor selections were limited mainly to those receptors (species) that are found in

the areas where Spartina is distributed, and to surrogate species for which sufficient life history 

and/or toxicological information existed so that reasonable exposure factors could be used to

estimate exposure and risk. The following bullets briefly summarize the ecological receptor guilds

for which exposure calculations were evaluated. Life history characteristics of these receptors are 

described fully in Chapter 4. 

• Mallard Duck (Anas platyrhynchus). This avian species was considered a representative 

(primarily herbivorous) waterfowl species bird that is common to the areas of interest. This 

species was evaluated due to its direct and indirect exposure through the consumption of 

aquatic plants. 

• Scaup (Aythya sp. [marila = greater; affines = lesser]). These species are more omnivorous

than the mallard, consuming a high proportion of their diet as animal protein, especially during 

spring and fall migration periods. Animal sources in the diet include mussels, small fish, and 

other benthic and pelagic invertebrates. The lesser scaup is considerably more common in 

Washington, but both are coastal species. 

• Red Fox (Vulpes vulpes). This mammalian species is a medium-sized primarily carnivorous

mammal of the canine family that is resident to much of western Washington, and whose range

is expanding. It is a surrogate for other carnivorous species such as the wolf, coyote, and 

mustellids. 

• Norway Rat (Rattus norvegicus). A mammalian species of near ubiquitous distribution in

lowland areas throughout Washington State and the U.S.. It is particulalry common around 

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coastal areas where it has been introduced through trade vectors of shipping. Rats are

commonly used for toxicity testing. 

• Deer Mouse (Peromyscus manisculatus). A common herbivorous mammal species found in

a variety of ecosystems, including coastal grasslands. Mice are also commonly used in toxicity

testing. 

• Bobwhite Quail (Colinus virginianus) A common avian test species. This species is not 

found in Washington State, but serves as a good surrogate for the introduced California (valley)

quail, Lophortyx californica, which is relatively common in western Washington. This species 

is primarily herbivorous, eating mostly seeds. Quail are commonly used to test avian 

sensitivity to toxicants. 

• Marsh Wren (Cistothorus palustris). A common native avian species to coastal grasslands 

and salt marsh habitats in Washington State. This species consumes a high proportion of its

diet in animal protein. 

• Cottontail Rabbit (Sylvilagus sp). A strictly herbivorous species common to much of 

western Washington, but introduced originally from the east coast. It is also a typical EPA test

species used particularly to evaluate dermal sensitivity. 

Threatened and Endangered Species and Species of Concern 

Threatened and endangered species are those species that have been given special legal and

protective designations by federal or state government resource agencies. A federally endangered 

species is one that is in danger of extinction throughout all or a significant portion of its range. A

federally threatened species is one likely to become an endangered species within the foreseeable 

future throughout all or a significant portion of its range. A species of concern is one for which status

information suggests the species is not abundant, and for which additional informatin is sought. 

Addressing exposure and risk to threatened and endangered species generally requires the use of

surrogate receptor guilds because they are rarely used (for obvious reasons) to establish toxicity

information on new chemicals (Sappington et al. 2000). A summary of all federal T&E species in 

Washington State, their Washington State status, and their potential for existence in areas where 

imazapyr treatments of Spartina could occur based on their habitat preferences is included in this

report as Appendix E. From this information, the potential exposure of T&E species is truncated to

only a few select species. In brief, utilization of Washington’s coastal areas by threatened and 

endangered (T&E) species is primarily limited to the listed salmonid species from the Columbia 

and Puget Sound basins. In addition, several coastal avian species listed as sensitive, candidate, or

state-monitor species are common to Willapa Bay and other areas where Spartina is distributed. 

2.3 Conceptual Model 

A conceptual model was developed for imazapyr (Figures 2-3 and 2-4). This conceptual model

accounted for the sources, pathways, and routes of exposure to the different trophic levels and 

ecological receptors. Exposure of ecological receptors to imazapyr used to control Spartina spp. 

could occur directly or inadvertantly (indirectly) through ingestion of contaminated food, water, or

sediment, through inhalation of aerosol, or through direct contact (e.g., insects). 

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Figure 2.3: Conceptual site model for imazapyr stressor impacts to terrestrial 

receptors.

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October 2003 Page 21
Figure 2.4: Conceptual site model for Imazapyr stressor impact to aquatic 

receptors.

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3.0 HAZARD ASSESSMENT 

The hazard assessment portion of a risk assessment summarizes environmental fate and toxicity 

data developed on the compound(s) of interest. To understand chemical hazards, some

fundamental aspects of toxicology should be clarified. A central tenet of toxicology is that there is 

some exposure dose at which no effect is measurable in the response tested, and this paradigm is

considered a valid model for this assessment. This dose or concentration is known as the no 

observable effect level or concentration (NOEL or NOEC). The lowest observable effect level or 

LOEL corresponds to the lowest dose at which a statistically significant difference is measurable

relative to an unexposed control group. Beyond these typical measures, standard toxicological

terms include the LC50, the exposure concentration that kills 50% of the animals tested; the EC50, 

the concentration that elicits a non-lethal effect in 50% of the organisms tested with the 

measurement endpoint. Measures such as the LC90 or EC90 simply reference variations in the 

proportion of the population tested that responds to the test (in this case, 90%). Other terms such

as the IC50 or IC10, reference a concentration that results in inhibition of an endpoint—in this case

50% and 10% inhibition respectively. These terms are often used to gauge the effect of a chemical 

on endpoints such as growth, or in-vitro endpoints such as the inhibition of an enzyme. 

In considering the hazards of imazapyr, it is important to recognize how the chemical can first

enter into commercial use. Imazapyr has been produced under different commercial formulations

with technical imazapyr and imazapyr isopropylamine salt (49 percent water solution) (Table 3-1). 

The formulations have the same mechanism of action on target plants, but different environmental 

factors control the efficacy of each formulation and where they might be applied. Most testing

related to the toxicity of imazapyr is related to the technical compound instead of the commercial 

formulation. The Arsenal® formulation listed below is the formulation projected for use in the 

estuary setting for Spartina control. Mechanism of action, environmental fate, and toxicity studies 

described in detail below reference these general formulations. 

Table 3-1: Product formulations of imazapyr.

Commercial Product 

% Imazapyr

Technical 

% Imazapyr

Isopropylamine salt % Inert Ingredient Source 

Arsenal

® N/A 25 75 other inert ingredients Cyanamid 1997 

Chopper® N/A 1 99 other inert ingredients Cyanamid 1997 

Arsenal

® Herbicide 28.7 N/A 71.3 other inert ingredients BPA 2000 

Arsenal

® Railroad 

Herbicide 

27.6 N/A 72.4 other inert ingredients BPA 2000,

USDA 1995 

Arsenal

® Applicators 

Concentrate Herbicide 

53.1 N/A 46.9 other inert ingredients BPA 2000 

Chopper® Herbicide 27.6 N/A 72.4 other inert ingredients BPA 2000 

Chopper® 22.6 5.4 72 other inert ingredients USDA 1995 

Chopper® RTU N/A 3.6 30 propylene glycol 

5.0 isopropanol 

61.4 other inert ingredients 

USDA 1995 

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3.1 Mechanism of Action and Efficacy

3.1.1 Mechanism of Action

The mechanism of action of an herbicide is the biochemical and/or physical method by which it

has been engineered to kill or suppress the growth of specific plants. Imazapyr (2-[4,5-dihydro-4-

methyl-4-(1-methylethyl)-5-oxo-1H- imidazol-2-yl]-3pyridinecarboxylic acid) is the active 

ingredient in the commercially available formulations Arsenal®, Chopper®, Stalker®, Assault®, and 

Contain®. The herbicide is commonly produced in either a weak acid form or as an isopropylamine

salt (Figure 3-1). The isopropylamine salt form is generally dissolved in a solution containing 49%

water, and most commercial products sold are in this salt form. Imazapyr was first registered under

the commercial formulation of the isopropylamine salt Arsenal® in 1984 and again with the weak

acid formulation Chopper

® in 1993 (Cyanamid Ltd. 1997). 

Figure 3-1: Chemical structures of the two forms of imazapyr. 

The specificity of an herbicide for target vegetation varies by herbicide family. Imazapyr belongs

to the chemical family imidazolinone. The imidazolinones are non-selective herbicides used to

control weeds, broadleaved herbs, and woody species. Imazapyr is primarily adsorbed through

plant tissue, but can also be adsorbed through roots in the soil. The compound is translocated in 

the xylem and phloem to the meristematic tissues. The mechanism of action is through inhibition 

of branched-chain amino acid synthesis. Specifically, imazapyr inhibits the enzyme acetohydroxy 

acid synthase (AHAS) or acetolactate synthase (ALS) which catalyzes the production of three

branched-chain aliphatic amino acids (valine, leucine, and isoleucine) that controls protein 

synthesis and cell growth (Cox 1996). 

Imazapyr is slow-acting and is generally most effective during post emergence axillary budding 

(Hanlon and Langeland 2000). Plants stop growth initially in the roots and continue in the above 

ground portions, with complete death occurring approximately one month after treatment,

depending on environmental conditions (Cox 1996). 

Animals do not synthesize their own three branched-chain aliphatic amino acids, but obtain them

by eating plants and other animals; therefore the engineered mechanism for plant toxicity is not

generally relevant to birds, mammals, fish or invertebrates. Toxicity associated with excessive 

doses administered to animals occurs by different mechanisms. 

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3.1.2 Efficacy on Target Vegetation 

Non-Spartina Studies 

Kay (1995) examined the efficacy of wipe-on applications of imazapyr on the common reed

(Phragmites australis) in aquatic systems. Similar to Spartina in the Pacific Northwest, the 

common reed is considered highly invasive in the midwest and Atlantic states, as well as 

Washington, serving little habitat value for fish and wildlife. In this field-based experiment, both

imazapyr (Arsenaltm) and glyphosate (Rodeotm) were tested at 0.5 and 0.25-strength dilutions with 

1% surfactant (X-77tm) added to the application mixtures. Control and cut plots received no

herbicide treatment. When examined at the end of the growing season, Kay observed significantly 

improved Phragmites suppression with Arsenal tm relative to Rodeotm, at both treatment

concentrations (Table 3-2). However, the overall control for either treatment chemical using the 

wipe-on application method was not considered acceptable. Short plants were shrouded from wipe 

treatment by larger plants, rendering complete control impossible. In addition, effects on nontarget emergent plants were documented. 

Table 3-2: Efficacy of Wipe-on Applications of Imazapyr and Glyphosate.

(source: Kay 1995)

1 

Treatment Rate % Dead in 1991

% Surface 

Covered with Live 

Reeds in 1992 

% Surface 

Covered with Live 

Reeds in 1993 

Control—No Herbicide or 

Mechanical Control 

0 3a 80a 78a

Cut Treatment—No Herbicide 0 100e 2d 4c

Arsenal 25% 57c 55b 64a

Arsenal 50% 75d 16c 55a

Rodeo 25% 38b 65ab 70a

Rodeo 50% 33b 47b 70a

Rodeo (spray) 1.25% 100e <1d 15b

1: Data in columns represent the mean of three replicates. Within column comparisons are not significant if the letter following

the value listed in a cell is found in another cell within the same column. Estimates of kill and survival were visually based. 

In a study looking at the response of torpedo grass (Panicum repens) to different application rates of 

imazapyr and imazapyr with fluridone, the level of control observed in the different plots varied

spatially and substantially depending on conditions where it was applied (Hanlon and Langeland 

2000). In this study, imazapyr (ArsenalTM) was applied in three canal systems of Lake Okeechobee at 

a rate of 0.28, 0.56, 0.84 or 1.12 kg acid equivalents (ae)/ha in a total tank mix volume of 187 L/ha 

(20 gal/acre) that contained 0.5% nonionic surfactant (Kenetictm). Some treatment plots also had the

imazapyr combined with 0.43 kg fluridone/ha. In one canal system where applied, highly effective 

control was obtained with imazapyr on one side of the canal, yet little or no control was obtained on 

the opposite side of the canal, despite similar treatment volumes and conditions. The authors 

speculated that floating periphyton mats in abundance on one side of the canal reduced the stem 

density of torpedo grass there and also may have bound up the applied herbicide such that the

torpedo grass along that side of the canal received a lower dose. Other key factors that may have 

affected the results of this study included: (1) soil moisture affected by hydroperiod fluctuations 

(from dry to 1.7 m inundation depth between areas), and (2) canopy of emergent thatch reducing the 

regrowth of the species is enhanced. Unfortunately, the authors did not attempt to analyze tissue 

concentrations in either target or non-target vegetation, or in the water such that differentiating the 

causes of the treatment differences could be resolved with less uncertainty. Notwithstanding, the

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study demonstrated clear control with imazapyr, particularly when the thatch overburden was burned

before herbicide treatment, as was done in one of the other canals where treatments were applied. 

When the thatch was not removed or burned, as was done in the third canal, no effective plant 

suppression was obtained. The addition of fluridone to the mixture did not appear to influence 

efficacy at any of the treatment concentrations. Although the results of these studies show that 

environmental conditions can highly affect the efficacy of imazapyr treatments in the aquatic 

environment, it could be concluded that predicting efficacy is far from a stochastic (random) process 

provided a basic understanding of the application conditions is considered prior to treatment. 

Table 3-3 summarizes some of the target aquatic and terrestrial nuisance plant species for which 

imazapyr has had reported efficacy (USACE 2003). 

Table 3-3: Examples of Aquatic Species Effectively Controlled by Imazapyr. 

(Source ACOE 2003)

Common Name Scientific Name

Giant Reed Arundo donax L.

Buttonbush Cephalanthus occidentalis

Purple Loosestrife Lythrum salicaria L. 

Melaleuca Melaleuca quinquenervia (Cav.) Blake 

Torpedo Grass Panicum repens L. 

Common Reed Phragmites australis (Cav.) Trin. ex Steud. 

Brazilian Peppertree Schinus terebinthifolius Raddi 

Giant Foxtail Setaria magna Griseb. 

Tamarisk or Salt Cedar Tamarix spp. 

Cattails Typha spp.

Para Grass Urochloa mutica (Forsk.) T.Q. Nguyen

There are a number of weeds that have developed resistance to imazapyr (Table 3-4). It is 

suspected that these plants have developed cross-resistance to imazapyr following the use of 

herbicides with the same mode of action (i.e., acetolactate synthase inhibition), primarily the 

sulfonylurea herbicides (Cox 1996). Resistance is afforded plants by developing a structurally 

similar form of the enzyme acetolactase synthase that is not blocked by imazapyr to the same

degree. The resistant enzyme form is thought to have developed primarily from a single point

mutation (Sathasivan et al. 1991). 

Table 3-4: Plants known to be resistant to the use of imazapyr. 

(source Cox 1996)

Common Name Scientific Name

Rigid ryegrass Lolium rigidum 

Kochia Kochia scoparia

Common chickweed Stellaria media 

Russian thistle Salsola iberica 

Perennial ryegrass Lolium perenne

Annual sowthistle Sonchus oleraceus 

Brassicaceae spp. Arabidopsis thaliana 

Algae spp. Chlorella emersonii 

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Spartina Control Studies 

Comparison studies of the efficacy of imazapyr relative to glyphosate on S. alterniflora control

have been recently conducted by Patten and Stenvall (2002) and Patten 2002. In these studies each

herbicide was tested at different application rates and/or drying times before tidal submergence or

rain. Comparisons were made in treated Spartina meadows at application rates of 0.84 and 1.68 kg 

ae/ha for imazapyr and 4.2, 8.4, and 18 kg-ae/ha for glyphosate. Imazapyr was dissolved in a cropoil surfactant while glyphosate was dissolved in a non-ionic surfactant at 1% v/v. The herbicides

were applied using a backpack sprayer equipped with a 1.5 or 3m boom equipped with Teejet 

11001 or 11002 nozzles that dispersed the herbicide mixtures at a rate of 94 to 940 L/ha. Efficacy

was evaluated based on a visual rating of percent control in a treated plot assessed approximately

one year following treatment compared to an untreated plot. The results of this study, representing 

numerous trials between 1997 and 2000, are reflected in the box plot in Figure 3-2 on the following

page. 

As demonstrated in Figure 3-2, a treatment concentration of 0.84 kg-ae/ha of imazapyr was slightly 

more effective than 8.4 kg/ha glyphosate for control of Spartina, but both treatments at these rates 

had high variability and produced incomplete control. At the higher rates of application of each 

herbicide nearly complete control was obtained (generally greater than 90%), and imazapyr had 

less variability around the median and slightly greater effectiveness at Spartina control than 

glyphosate. As depicted, more than 10-times the amount of glyphosate was required to achieve

similar control as was obtained with imazapyr, but imazapyr treatments produced a greater number

of outliers at the higher treatment rate. The variability in treatment response for either herbicide

was not discussed by the authors. Treatment variability may have been due to factors such as

overlying thatch (Hanlon and Langeland 2000), differences in plot size between years, or

surfactant differences. 

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Figure 3-2: Box and whisker graph of Spartina control as a function of rate of 

imazapyr and glyphosate use in experiments conducted in Willapa Bay, WA from 

1997 to 2000. 

(Source: Patten and Stenvall 2002.)

Note: Boxes in box-plots depict the data between the 25th and 75th percentiles, with the cross 

bar in the box representing the median of the data, and the hinges (whiskers) representing the

highest and lowest data values for the main body of data (5th and 95th percentiles); dots beyond 

the whiskers are considered outliers by standard statistical convention. 

Effects of drying time were examined in a separate experiment using the 1.68 kg/ha imazapyr

treatment and 8.4 kg ae/ha glyphosate treatment only (Patten and Stenvall 2002). Drying times for

this experiment were segregated by 4-7 hours and greater than 7 hours. Patten and Stenvall’s study 

found that imazapyr provided more control with less variability and shorter dry time requirements

when compared to glyphosate (Figure 3-3). Glyphosate at 8.4 kg/ha was not as effective at

Spartina control when provided a drying time of only 4-7 hours. Even when allowed at least seven 

hours of dry time, the glyphosate treatment exhibited greater variability around the median than 

either of the imazapyr dry-time treatments. In contrast, imazapyr efficacy was not significantly

different between the 4-7 and > 7 hour dry times investigated. While the results of this experiment 

demonstrate the sensitivity of glyphosate to drying time, the results would have been more useful if 

the glyphosate treatment concentration that provided nearly complete control (18 kg ae/ha) was 

compared against the imazapyr treatment that provided similar Figure 3-3). The comparison 

depicted in Figure 3-3 leaves in question what factor dose may have played in this efficacy trial. 

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Figure 3-3: Box and whisker graph of Spartina control as a function of dry time for 

imazapyr (1.68 kg/ha) and glyphosate (8.4 kg/ha) use in experiments conducted in 

Willapa Bay, WA form 1997 to 2000. 

(Source: Patten and Stenvall 2002.)

Additional studies by Patten (2002) examined the efficacy of imazapyr and glyphosate with

various seasonal timings, spray volumes and with several surfactants. Formulations of these 

surfactants are proprietary and therefore a full analysis of their components is not possible (their 

toxicity is discussed in Section 3.5). Seasonal application timings corresponded to the stage of 

plant development from preanthesis (late June to mid-July), anthesis (mid-July to August), or seed

filling (September). Application volumes ranged from 23 to 374 L/ha at active ingredient

concentrations of 0.84 and 1.68 kg-ae/ha for imazapyr, and 7.2 and 8.4 kg/ha for glyphosate and 

application volumes from 94 to 748 L/ha at concentrations of 3.63 to 8.4 kg/ha for glyphosate.

Surfactants evaluated included R11tm, Agri-Dextm, Hastentm, LI100tm, Syl-Tactm, Kinetictm, and

Dyne-Amictm. 

The efficacy of imazapyr and glyphosate relative to application date were inconsistent, and shown 

to be more influenced by spray volume than by application timing, provided application occurred

during the growing season. Higher spray volumes tended to result in more consistent Spartina

control; however, this trend was not consistent across all dates and sites (Table 3-5). This

inconsistency was considered due more to changes in estuarine conditions (e.g., storms, tides, mudcovered leaves) than to physiological changes in Spartina during different seasons. The least

effective dates for imazapyr efficacy were early July and October. 

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Table 3-5: Effect of rate, timing and spray volume in 2000 on the efficacy of imazapyr and 

glyphosate for smooth cordgrass control in Willapa Bay, WA. 

(Source: Patten, 2002.)

Application

Spray

Treatment Efficacy

% Control 

Herbicide 

Rate 

kg/ha 

Application Date in

Year 2000 Volume L/ha Site 1 Site 2 

Imazapyr 0.84 July 6 94 40 35

August 2 94 81

August 31 94 47 89

September 14 94 63

October 7 94 19 90

July 6 468 44

August 2 468 49

August 31 468 92

September 14 468 84

October 7 468 23 46

Imazapyr 1.68 July 6 94 83

August 2 94 94 99

August 31 94 36

September 14 94 94 96

October 7 94 44

July 6 468 77

August 2 468 97

August 31 468 92

September 14 468 93

October 7 468 59

Glyphosate 8.4 July 6 94 20 0

August 2 94 30

August 31 94 0 69

September 14 94 32

October 7 94 62 46

Untreated 0 0

Imazapyr at 1.68 kg/ha provided excellent control of Spartina when applied at ultra-low application volumes (23 and 47 L/ha) (Table 3-6). The study showed that as long as drying time was

sufficient, good control could be achieved with low volume applications. Across all sites and 

experiments described in Patten’s 2002 paper, 58 and 88% cordgrass control was obtained with

imazapyr applications at 0.84 and 1.68 kg ae/ha, respectively. Glyphosate, by comparison, 

provided 45 and 81% control, which was obtained at the 7.2 and 8.4 kg/ha application rates, 

respectively. All surfactants used in the study provided delivery of the herbicides to yield effective 

control of the target organism. 

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Table 3-6: Effects of herbicide rate and application spray volume in August 1999 on

smooth cordgrass control in Willapa Bay, WA.

(Source: Patten, 2002,)

Rate

Spray

Volume 

Treatment efficacy 

% control 

Herbicide kg/ha L/ha Site 1 Site 2 Site 3 Site 4 Site 5 Site 6 

Imazapyr 1.68 23 93 93

Imazapyr 1.68 47 96 99 93 84 96 95

Glyphosate 8.4 94 85 52 89 48

Untreated 0 0 0 0 0 0

3.2 Environmental Fate and Chemistry 

3.2.1 Physical Chemistry 

Solubility 

Imazapyr is ionized under typical environmental conditions of pH 5-9, and is therefore highly 

soluble in water. The solubility of imazapyr varies somewhat with the product formulation. The 

solubility of imazapyr increases with temperature. For example, the solubility of the compound is

reported as 9,740, 11,272, and 13,470 mg/L (ppm) at 15 o

C, 25 oC, and 35 oC, respectively 

(Mangels and Ritter 2000). Typical temperatures of application in Washington State would 

bracket the solubility measures recorded between 15 o

C and 25 oC. Generally, solubility will 

bracket the range of 1 to 1.5% (i.e., 10,000 to 15,000 mg/L) in water at 25 o

C. A saturated 1%

solution of imazapyr in freshwater at 25 o

C will exhibit a pH of approximately 3 to 3.5 (Toxnet

2003). Because of the high solubility of the compound, it has inherently low sorption potential, 

and relatively high potential for mobility through soils. 

The octanol:water partition coefficient (LogP) of imazapyr is reported as 1.3, reflecting its high

solubility in water, low solubility in lipid (octanol), and hence low propensity to bioconcentrate or 

bioaccumulate (ToxNet 2003). 

Imazapyr has a melting point range of 169 to 173 o

C. 

Dissociation constants reported for imazapyr reflect its ionization potential under typical

environmental conditions. For reader clarification, the pH at which an acid is 50 percent 

dissociated between its non-ionized and ionized forms is called its pKa. Thus, when the pH of a

solution is equal to its pKa the chemical will be dispersed equally between an ionized and 

unionized state. Imazapyr dissociates at two different pH levels, with dissocation constants (pKa)

of 1.9 and 3.6. In general, ionized forms of chemicals represent lower ecological risk because they 

are unable to penetrate cell membranes due to low lipid solubility. For acids such as impazapyr, as

the pH is elevated above the pKa the proportion of the compound in an ionized state will increase.

In the marine intertidal mudflats where the imazapyr would be applied to control Spartina, the pH 

of sediment surfaces and sediment pore water should be elevated above neutral, and the compound 

will be entirely in an ionized state (Figure 3-4). However, surfactants applied with the product are 

designed to facilitate uptake for product efficacy, and therefore reduce this element of protection. 

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Figure 3-4: Imazapyr Dissociation Under Different pH Conditions. 

(Source: Mangels and Ritter 2000.)

3.2.2 Environmental Degradation 

Synopsis 

The rate and form of degradation of imazapyr varies somewhat with the environment where it is

applied. Movement within the environment (e.g., through soils, water, plants, sediments) of the 

weak acid is primarily determined through by the pH of the host system. The primary form of

degradation in water is via photodegradation. Photolysis half-lives in water have been reported at

2 days; however degradation decreases with increasing pH. Ozone may also degrade imazapyr

when applied in a water treatment setting (Rashin and Graber 1993). Imazapyr has been detected

in surface and ground water samples taken by the Washington State Department of Ecology 

following aerial application on forest lands, although regular monitoring of the herbicide is lacking 

(Cox 1996). In soils, degradation is primarily driven by microbial metabolism. Microbial 

metabolism in sediments has not been thoroughly investigated. 

Soil Adsorption and Degradation

Imazapyr will adsorb to soils and sediment weakly. It has a reported organic carbon partition 

coefficient (Koc) in soil of 142 cc/g (Mangels and Ritter 2000). Thus, imazapyr is considered

relatively mobile in soils. Adsorption is pH dependent, again, reflecting its propensity for

ionization at pH levels above its pKa (Figure 3-4). For example, a pH below 5.0 in soil limits

movement due to an increased adsorption capacity within the soil; whereas above a pH of 5.0 

concentrations of imazapyr become negatively charged (i.e., ionized) and do not bind well with

soils, which increases its mobility. The adsorption coefficient for Arsenaltm, an isopropylamine

salt formulation, varies for different types of soil. The adsorption coefficient reported for clay

loam soils is 1.7 verses 4.9 for silt loam soils with 4.0 percent organic matter (PMEP 1985).

Leaching has been observed up to 50 cm in soil. Another study related to imazapyr mobility in soil 

observed significant residues to a depth of 1.5 to 3 m (4.9 to 9.9 feet), depending on application 

rate (Cox 1996). Leaching has been observed up to 50 cm in soil. Another study related to

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October 2003 Page 32
imazapyr mobility in soil observed significant residues to a depth of 1.5 to 3 m (4.9 to 9.9 feet), 

depending on application rate (Cox 1996). 

The primary degradation pathway for imazapyr in soils is through microbial metabolism, with

photolysis and other degradation pathways providing a limited source of degradation. During

aerobic microbial metabolism, the imidazolinone ring is opened and a hydroxy metabolite is

formed as a result of the conversion of the carboxilic acid group on the pyridine ring (Figure 3-5). 

Reported half-lives of imazapyr (technical grade) in soil range from 25-141 days (Cyanamid Ltd. 

1997, Cox 1996). Most of the reported variation in soil half-life is related to detection method 

applied in the study: plant injury versus laboratory analysis. 

Figure 3-5: Degradation Pathway of Imazapyr. 

(Source: Mangels and Ritter 2000.)

Recent studies by the manufacturer examined the aerobic metabolism of imazapyr in sandy loam

soils (Mangels and Ritter 2000). They proposed a half-life of 117 days based on degradation 

through the first 28 days of the study. However, after 4 months, when the study concluded, only 

26% of the dose applied had degraded, and only 5.6% of the dose had completely mineralized

(degraded) to CO2 . The authors considered the rapid early degradation to reflect the more active

microbial community in the soil initially available, as the soils used in the testing were derived 

from an agricultural field (Figure 3-4). In a second, 12-month study also using sandy loam soil

with an application rate of 1.0 to 1.5 ppm, 66% of the applied dose still remained at the end of the 

study. No volatilization of the parent compound occurred. When using the data from 0 to 9 months 

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only, the half-life was estimated at 12.8 months, but the final three months of the study revealed no 

significant additional degradation, extending the half-life estimate to 17 months. A replicate 12 

month study showed 88% of the applied imazapyr was recoverable after 365 days. These

collective results reveal that degradation rates can vary substantially even within an experiment. 

Figure 3-6: Degradation Rate of Imazapyr in Sandy-Loam Soil. 

(Source: Mangels and Ritter 2000.)

The key parameters that appear to effect the soil degradation rate of imazapyr in the field include

temperature, organic carbon, and particle size. In a study by McDowell et al. (1996), the half-life

for imazapyr (from Arsenaltm) was addressed using a bioassay approach that gauged the time taken 

for the amount of herbicide in the “plant-available” pool to equal half of that which was applied.

Using the lentil growth bioassay in New Zealand Templeton silt loam soils, the half-lives varied by 

approximately 87 days depending on temperature, soil organic matter, and microbial biomass

carbon (μg-C/g soil), with a range of 68.6 to 155.4 days. Degradation of the technical grade

imazapyr in soils through microbial activity reportedly increased with the following environmental

conditions (1) warmer temperatures (15°C vs. 30°C), (2) presence of sandier soils (i.e., sandy loam

vs. clay loam), (3) aerobic soil conditions, (4) increasing soil moisture, (5) increasing pH, (6) lower 

organic matter soil. However, the effect of organic matter could not be isolated from the effect of 

pH under their experimental design, so the impact of these variables on degradation rate was not

entirely conclusive. 

In another study by the manufacturers of imazapyr, Cyanamid Ltd. (1997), reported that the 

imazapyr isopropylamine salt degraded 6 months faster (3 verses 9 months) at 45°C than at 37°C. 

The related imazapyr acid formulation also degraded faster with the same conditions as the 

technical grade imazapyr. 

Anearobic soil metabolism of impazapyr appears to be insignificant from the existing studies

conducted to date. In one study, soil was treated with impazapyr at 1.0 ae/acre. 

There are also reports of imazapyr “leaking” out of the roots of treated plants and impacting 

surrounding native vegetation. A study by Lee et al. (1991) reported an increase of imazapyr

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residues 231 days after treatment due to runoff of residues from plant surfaces after rainfall and 

release from decaying plant matter. Rainfall after application of imazapyr has shown to increase 

the ability of the chemical to be adsorbed, however impacts due to increased mobility in the soil 

column may outweigh efficacy improvement. 

Degradation in Aquatic Environments 

Conditions in aquatic environments differ substantially from those of terrestrial soils, both in terms 

of the regular exchange of waters within the sediment (porewater) and over it, and in the range of 

oxygenation experienced in typical sediments that can affect microbial metabolism (i.e., aerobic

verses anaerobic ratios). Early studies in freshwater, primarily by the product registrant, examined

the pathways and mechanisms of degradation in water and underlying sediment (Mangels and 

Ritter 2000). Typical controlled degradation studies examine the rate of degradation by the 

accumulation of radiolabled CO2, which represents the final breakdown product of a radiolabled 

parent compound and its intermediate degradation products. These studies are done either in 

water, moist sediment, or in slurries of water and sediment. 

The degradation of imazapyr when applied directly to water largely mimics the pathway by which 

the herbicide would be solubilized at high tide after application to Spartina during low tide. 

Residual imazapyr on the plants that may not have completely dried or adsorbed will be inundated 

by the incoming tide and presumably solubilize. Aquatic degradation studies with imazapyr 

applied to a freshwater surface directly have shown that imazapyr initially photodegrades rapidly 

to two primary products, “CL 119060”, and “CL9140” (Figure 3-5). According to the 

manufacturers, CL119060 is biologically oxidized to CL 9140, and eventually mineralizes to 

carbon dioxide (CO2) following the cleavage of the pyridine ring structure. Hydrolysis of the 

parent compound was found to be negligible, with controlled experiments in distilled water,

documenting only 3% of hydrolysis product (CL252974—see Figure 3-5) accumulated after 12 

days of incubation in pH 9 water. This hydrolysis product gradually increased over a 30 day 

incubation period to 6.9% of the recovered product, but the pH 9 value as tested would rarely be

seen in the estuarine environment and hydrolytic mechanisms of degradation in situ would 

probably be less than observed in this study. 

In a controlled aerobic aquatic study with both photodegradation products applied at 0.083 ppm in 

Missouri and Florida pond water, less than 22% of the applied radioactive dose dissipated from the

water phase into the sediment phase. The quantity of CL 119060 decreased from 77 to 0.5% of the

administered dose in this replicated laboratory experiment while the concentration of CL 9140 

initially increased (Figure 3-7—Missouri data only). However, both imazapyr degradation 

products rapidly degraded, with half lives less than or equal to 3 days (Mangels and Ritter 2000). 

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Figure 3-7: Degradation of Imazapyr’s Initial Photodegradation Products CL 

119060 and CL 9140 in a controlled aerobic aquatic system using Missouri pond 

water.

(Source: Mangels and Ritter 2000.)

Unlike lab degradation experiments where more variables can be controlled and measured, field 

experiments are generally termed “dissipation” studies, as the multiple variables inherent to such 

systems limit the range of analyses that can be conducted. With this understanding, 

complementary field dissipation experiments were conducted by the product registrant in shallow

Florida and Louisiana freshwater pond systems where the parent product imazapyr (as Arsenaltm) 

was applied to the surface of the water at 1.5 lb ae/acre (883 ppb). Dissipation (field degradation)

was followed in water and sediment over 180 days. Figure 3-8 reflects study results from water

and sediment analyses from the Louisiana pond study through the first 30 days of study, over

which period the vast majority of dissipation had occurred. Similar results were obtained with the

Florida pond system (not shown) although degradation was slightly faster and there did not appear

to be the initial spike in the sediment concentration that was observed in the Louisiana pond system

(Figure 3-8). The first-order half-lives in the water and sediment were 1.9 and 12.8 days, 

respectively. No detectable residues of imazapyr were found in the water and sediment after 14 

and 59 days, respectively. 

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Figure 3-8: Residues of imazapyr in water and sediment from a Louisiana pond 

treated with 1.5 lb ae/acre. 

(Source: Mangels and Ritter 2000.)

Tidal flux in estuarine environments also provides a consistent and predictable rinsing effect that

will solubilize applied herbicide and contribute to its removal from an area where recently applied.

Freshwater degradation studies in slurries probably represent the most similar laboratory 

conditions for degradation comparisons to tidal environments. However, to our knowledge these 

tests have been done in freshwater only. These conditions challenge the researcher attempting to 

isolate the variety of mechanisms that could be responsible for degradation of imazapyr in an

estuarine setting. Recent studies have addressed some of the uncertainty. Patten and Stenvall 

(2002) examined the fate of imazapyr applied directly to sediment to gauge its persistence over 

time. In this study, imazapyr concentrations were measured in intertidal waters and sediments 

adjacent to a Spartina meadow. Analyses were conducted 3, >24 and > 48 hours after application 

at the standard treatment concentration of 1.68 kg ae/ha used in the efficacy trials discussed earlier.

Sediment samples collected three hours after application were retrieved immediately after the first 

tidal wash over the treated area. The study design was conservative in that there was no 

intercepting algal or emergent vegetation overlying the sediment where the herbicide was applied.

In this study, the maximum geometric mean concentration of imazapyr detected in sediments from

four replicated trials detected 3-hours application was 5.4 mg/kg-sediment. Twenty-four to

seventy-six hours after application the geometric mean maximum detection was 2.26 mg/kgsediment, roughly half of that detected after three hours. In the water, 0.119 mg/L imazapyr was 

detected after 3-hours, and less than 0.00006 mg/L (the method detection limit) was detected after 

24 to 48 hours. The intent of this study, however, was primarily geared towards addressing 

ecological risk under a conservative application scenario, as opposed to the in situ degradation. No 

“time zero” data were reported in the study results, nor were other environmental conditions that 

may have effected the results reported such as temperature, organic carbon, etc. Without such 

data, the in situ degradation can only be estimated. 

Some of the experimental design issues with the preceding experiment were addressed in a more

intensive fate study conducted by Patten subsequently (2003). In this study, imazapyr was again

applied to bare mud flat at 1.68 kg/ha with 1% (v/v) Agridex surfactant to a plot size of 30 x 33 m 

in the upper intertidal zone. The tidal front at this site could cover the application area in 13 

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minutes. A standard backpack sprayer with a 3 m boom was used to apply the herbicide/

surfactant mixture at a rate of 97L/ha in the early morning. The pH 7.9 sediment contained 49%

water, and 51% dry matter. The solid constituents of the sediment contained 5.4% organic matter,

and 18.3, 65.5, 16.2 percent sand, silt and clay, respectively. Water was collected in 1L jars buried

within 1 cm of their lips in the sediment to capture the incoming tidal front after application, and 

the jars were spaced in triplicate at 10 m intervals along the tidal front “grid”. Water samples were

collected after the 1st, 2nd, 3rd, and 7th tides, corresponding to 3.5, 14, 28 and 77 hours after

treatment. Square sediment cores 8-cm deep were obtained in similar positions along the grid as

the water samples, but unlike the previous experiment, samples were taken immediately after 

application (time 0), as well as the 1st, 2nd, 6th, 14th, 28th and 56th tidal exchange after treatment.

These exchanges corresponded with 1, 14, 27, 77, 184, 366 and 703 hours after the initial sediment 

application. Finally, sediment sub-samples were obtained from triplicate Spartina plots (3x4 m)

treated with the same application regime as the bare sediment to gauge canopy interception under 

typical treatment regimes. In this trial, the treated Spartina was 1.7 m tall, and was concentrated in

an area approximately 2 m above Mean Lower Low Water (MLLW) MLLW. All of these areas of 

treatment were then compared against samples taken from untreated bare sediments. 

Figures 3-9 and 3-10 reflect Patten’s (2003) results on the water and sediment persistence of

imazapyr. As depicted, measurable concentrations of imazapyr declined exponentially in both the 

water and sediment. Following applications to the bare intertidal mud flat, the maximum average

water concentration detected was 3.4 μg/L (ppb) and the maximum average sediment 

concentration detected was 5.4 mg/kg (ppm). The “zero asymptote” was approached at 40 and 400 

hours for the water and sediment, respectively (Figures 3-9 and 3-10). 

Figure 3-9: Persistence of imazapyr in estuarine waters of Willapa Bay following 

direct application to an unvegetated tidal mud flat. Data represented are mean 

values of triplicate samples +/- SE. 

(Source: Patten 2003.)

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Figure 3-10: Persistence of imazapyr in estuarine sediment in Willapa Bay, WA 

after direct application to an unvegetated tidal mud flat. Data represented are 

mean values of triplicate samples +/- SE. 

(Source: Patten 2003.) 

Persistence in Biological Tissues 

Biological tissues may act as an additional reservoir for chemicals applied intentionally or

inadvertently to the environment. When an organism accumulates chemicals in its tissues 

following direct exposure it is known as bioconcentration. If the organism is consumed (predated

upon) by another organism resulting in a higher concentration of the chemical in the predator, the

chemical is considered to bioaccumulate in the “food web”. In simple terms, the chemical

accumulates at a rate faster than normal metabolic processes eliminate it. Although 

bioconcentration and bioaccumulation may have toxicity implications, toxicity varies by chemical

and dose, thus these mechanisms should be considered independently when evaluating the 

biological fate of applied herbicides. As indicated in the discussion of physical chemistry (Section

3.1) the octanol:water partition coefficient and its high solubility indicate the compound is not apt 

to concentrate in tissue. Notwithstanding, to address imazapyr and its degradation products in 

biological tissues, several treatment studies of relevance have been conducted in Florida and 

Missouri pond systems that contained bluegill, tilapia, catfish and crayfish (Mangels and Ritter 

2000). The ponds contained 75, 28, 213 or 261 ppb imazapyr following treatments of Arsenaltm to

the banks and outer edges of the ponds at a rate of 1.6 lb ae/acre in spray solutions of 21 to 23 gal.

Ultimate concentrations in the ponds varied due to dilutional profiles inherent to the ponds (e.g., 

volumes). Table 3-7 summarizes the principal findings from this study in each of the pond systems

evaluated. 

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Table 3-7: Persistence and Bioconcentration of Imazapyr in Missouri and Florida Pond

Water, Sediment and Resident Aquatic Biota.

(Source: Mangels and Ritter 2000.)

Metric 

Measured 

Untreated 

Control 

Pond 

Pond 1 

(MO Pond

#11) 

Pond 2 

(Fl pond # 

11) 

Pond 3 

(MO Pond 

#21) 

Pond 4 

(FL Pond

#21) 

Initial Water 

Concentration

μg/L (ppb) 

< 0.207 28 75 213 261

Initial Sediment 

Concentration

μg/kg (ppb) 

<0.475 ~1.51 ~2.51 ~10.21 ~ 10.21

Half-life in Pond

Water (days)

Not applicable 14.1 8.4 14.5 3.9

Half-life in Pond

Sediments 

Not applicable Not calculated2 Not calculated, 

(too few data) 

Not calculated2 9.2 

Residue in 

bluegill

All samples 

<5.35 ppb 

< 50 ppb (MDL3) < 50 ppb (MDL3) 3 hrs post trt. =

0.636 ppm,

thereafter < 50 

ppb (MDL3), 

< 50 ppb (MDL3) 

Residue in 

catfish 

All samples 

<5.35 (except 1 

sample at 14.1) 

< 50 ppb (MDL3) < 50 ppb (MDL3) 3 hrs post trt. =

0.233 ppm,

thereafter < 50 

ppb (MDL3) 

< 50 ppb (MDL3) 

Residue in tilapia All samples < 

5.35 

< 50 ppb (MDL3) < 50 ppb (MDL3) 3 hrs post trt. =

0.068 ppm,

thereafter < 50

ppb (MDL3) 

< 50 ppb (MDL3) 

Residue in 

crayfish

All < 5.35 

(except 1 @ 

10.6) 

< 50 ppb (MDL3) < 50 ppb (MDL3) 3 hrs post trt. =

0.059 ppm,

thereafter < 50 

ppb (MDL3) 

< 50 ppb (MDL3) 

1: Based on interpretation of graphical data prepared by the researchers 

2: Sediment levels persisted in these ponds; the authors attributed this to an unusual inversion that resulted in

pond turbidity prior to pond treatment and reduced the rate of photodegradation 

3: MDL = method detection limit 

A separate study examined the potential for bioconcentration and persistence in a mollusc species, 

the freshwater clam (Corbicula fluminea). In this study, clams were exposed to the Arsenaltm

formulation of imazapyr in a mesocosm containing water and sediment, and the water was

inoculated with 0.091 mg ae/L. Similar to the study results reported above, no imazapyr was 

detected in the clam tissue at or above the 50 μg/kg (ppb) detection limit. Over the 28-day study,

the concentration of imazapyr in the water declined only minimally, from 81 to 75.1 ppb, while the

sediment concentration increased from non-detectable to 29.2 ppb at the end of the experiment.

No toxicity was reported. 

3.3 Toxicity to Terrestrial Receptors 

In this section we examine data from acute, sub-chronic, and chronic imazapyr exposure studies

with terrestrial wildlife and invertebrates. The toxicity of imazapyr to these ecological receptors is

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discussed, as known, and data gaps are identified. The descriptions of toxicity in Table 3-8 

established by the Environmental Protection Agency (EPA) are used as a template to qualitatively 

gauge study results discussed. As depicted, these descriptions vary slightly depending on the class 

of animal being tested (avian or mammalian) and the study being performed (acute oral LD50 or

dietary LC50). The LD50 is the statistical derivation of a dietary or drinking water dose, which is 

predicted to cause 50% mortality in the given population being tested. The LC50 is a similar 

number, based on the concentration of a compound in air or water. The criteria for these 

descriptions are presented below in Table 3-8. The specific toxicity data discussed below are

separated into simple animal classifications (e.g., mammals, birds, insects, etc.) and the studies

conducted generally reflect EPA protocols with standard test species. In using this toxicity

classification scheme it becomes possible to qualitatively compare toxicity values of the active 

ingredient and product formulations amongst species. 

Table 3-8: Hazard Classifications to Address Wildlife Risk from Herbicide Use. 

(EPA 1995)

Mammals Mammals Avian Avian

Hazard Category 

Acute Oral or Dermal LD50

(mg/kg) 

Acute Inhalation LC50 

(ppm) 

Acute Oral LD50

(mg/kg) 

Acute Inhalation

LC50 (ppm) 

Very highly toxic <10 <50 <10 <50 

Highly toxic 10-50 51-500 10-50 50-500 

Moderately toxic 51-500 501-1000 51-500 501-1,000 

Slightly toxic 501-2,000 1001-5000 501-2,000 1,001-5,000 

Practically non-toxic >2,000 >5,000 >2,000 >5,000 

It must be recognized that species differences in terrestrial ecological receptors (mammals, birds,

etc.) may exist that are not predictable from the classification scheme represented in Table 3-9. 

Some wildlife receptors that may be at risk of exposure to imazapyr are rarely used in toxicity 

testing for lack of a consistent supply and approved protocols. For example, we found few data on 

omnivorous and carnivorous species such as the raccoon and coyote, large ungulates such as the

black-tailed deer that commonly forage along estuary margins, and migratory shorebirds, 

passerines, and reptiles and amphibians. 

The use of surrogate test species with similar dietary and/or behavior patterns has been shown to

provide a relatively reliable predictor of toxicity for the most sensitive species of fish (Sappington

et al. 2000). A similar relationship likely exists for other wildlife receptors that are not routinely 

used for toxicity testing when compared against surrogates. However, only site-specific risk

assessments would be able to fully quantify risks to resident and migratory wildlife receptors from 

chemical exposure in each location where Spartina control with imazapyr is envisioned. This

assessment therefore must use surrogate species such as the rat and rabbit to gauge toxicity to other

wildlife that may be more likely to be found using the habitat found along the state-managed 

roadways of Washington State. The rat provides a reasonable surrogate of an omnivore, the rabbit

an exclusive herbivore, and the quail and duck provide surrogates of upland and wetland bird

species, respectively. 

3.3.1 Mammals 

Based on EPA criteria specified in Table 3-9, imazapyr would be considered practically non-toxic to 

mammals based on acute and chronic studies conducted with a variety of mammalian species. For 

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example, the reported acute oral LD50 concentration for rats is >5,000 mg/kg. No significant 

bioaccumulation has been reported in mammals. Rats were observed to rapidly excrete imazapyr in

urine and feces with no residues detected in their liver, kidney, muscle, fat, or blood. Results from a 

series of tests looking at the mammalian response to acute oral, dermal, and inhalation

administrations of imazapyr isopropylamine technical, imazapyr technical, and imazapyr

isopropylamine was compiled by Cyanamid Ltd. (1997) (Table 3-9). 

Table 3-9: Acute and Subchronic Mammalian Toxicicity to Imazapyr. 

Test 

Substance 

Animal 

Species 

Administration 

Route Gender 

LD50 or 

ED50 body 

wt., (or) 

ppm diet* Effect 

Testing Facility,

(reporting year)

Male > 10,000* DA Oral

Female > 10,000* DA

Male DA, B, A, S, CY,

C, DBW Intraperitoneal 

Female 3,700 DA, B, A, S, CY,

C, DBW 

Male > 5,000 DA Subcutaneous Female > 5,000 DA

Male > 2,000 NOEL

Rat 

Dermal Female > 2,000 NOEL

Male > 10,000 DA Oral Female > 10,000 DA

Male 3,450 DA, B, A, S, CY,

C, DBW Intraperitoneal 

Female 3,000 DA, B, A, S, CY,

C, DBW 

Male > 5,000 DA, B, S 

Imazapyr 

isopropylamine 

technical 

(49.3%) 

Mouse 

Subcutaneous Female > 5,000 DA, B, S 

Medical Scientific

Research, 

Laboratory (1983) 

Male > 5,000 NOEL Rat Oral

Female > 5,000 NOEL

Male > 2,000 NOEL Rabbit Dermal Female > 2,000 NOEL

American Cyanamid

Company (1983)

Male > 1. ND

Imazapyr 

technical

Rat Inhalation Female > 1.

(analytical) ND 

Food and Drug 

Research 

Laboratories (1983) 

Male > 5,000 DA Rat Oral

Female > 5,000 DA

American Cyanamid

Company (1983)

Male > 5,000 DA Mouse Oral Female > 5,000 DA

American Cyanamid

Company (1986)

Male > 2,148 NOEL Rabbit Dermal Female > 2,148 NOEL

American Cyanamid

Company (1983)

Male > 0.2 NOEL 

Imazapyr 

isoproplyamine 

25% AS 

Rat Inhalation Female > 0.2 

(analytical) NOEL 

Food and Drug 

Research 

Laboratories (1983) 

NOEL = no toxic signs, DA = decreased activity, ND = nasal discharge, B = blepharoptosis, A = ataxia, S = sedation, CY = 

cyanosis, C = convulsion, DBW = decreased body weight 

Source: Cyanamid Ltd. (1997) 

4,200

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Significant sub-lethal effects were reported for each formulation of imazapyr tested at doses that

exceeded the “practically non-toxic” acute lethal criteria for administration routes, except the

inhalation route, where sublethal effects occurred at lower doses (Table 3-9). The most significant

effect was found with technical grade imazapyr isopropylamine administered via intraperitoneal 

injection, although because the other forms of imazapyr did not test reactions using this method, it

is hard to draw conclusions on the relative toxicity of imazapyr isopropylamine technical grade.

Furthermore, this method of toxicant administration is not environmentally relevant because

mammals would not be dosed in this manner in a natural setting. No overall observable effect was

noted for any formulation of imazapyr administered dermally, and effects in mammals exposed to 

imazapyr via inhalation were only observed with imazapyr technical grade. 

Technical grade imazapyr is reported as moderately irritating to rabbit eyes, with complete 

recovery within 7 days of exposure (BPA 2000, Cyanamid Ltd. 1997). The same result was found 

with imazapyr isopropylamine 25% AS. Imazapyr technical is also reported as mildly irritating to

rabbit skin (BPA 2000, Cyanamid Ltd. 1997). Studies reviewed by the EPA concluded that

imazapyr technical is corrosive to the eyes and can cause irreversible eye damage (Cox 1996).

Commercial formulations of imazapyr appear to be less toxic by this route of exposure. 

Dermal exposure studies have shown imazapyr to yield statistically inconsistent sublethal effects. 

Where an effect was observed, it was usually observed as erythema, a localized increase in blood 

flow observed as a ‘reddening’ or rash-like symptom. A 21-day sub-acute rabbit dermal toxicity 

study at doses of 0, 100, 200, or 400 mg/kg/day revealed no consistent pattern of toxicity, and the

NOEL was concluded to be the highest dose tested (HDT) (400 mg/kg/day) (Fed Reg, 62, 1997)

even though a higher dose would most likely be more accurate. Another study with imazapyr

isopropylamine (25% AS) was reported to cause slight erythema at 24 and 72 hours post-dermal 

exposure in rabbits; however, the same formulation was observed to form no dermal reaction in

guinea pigs (Cyanamid Ltd. 1997). Other dermal exposure studies in both rats and rabbits report

LD50 concentrations at > 2,000 mg/kg, and > 2,148 mg/kg, respectively (Table 3-9). 

The product registrant conducted subchronic dietary toxicity tests where imazapyr isopropylamine

was administered orally at concentrations of 0, 1000, 5000 and 10,000 ppm for 13 consecutive 

weeks (Cyanamid Ltd. 1997). The study reported the maximum NOEL for rat diets as 5,000 ppm 

(325 mg/kg/day in males and 370 mg/kg/day in females). Another 90-day rat feeding study at

doses of 0, 15,000, and 20,000 mg/kg-diet yielded a NOEL values of 1,695 mg/kg/day the HDT, 

and an estimated (Fed Reg 1997). 

Chronic Testing

Chronic toxicity studies have been conducted with mice, rats, dogs and rabbits to address effects

on survival, carcinogenesis, teratogenecity and intergenerational effects. Dogs fed doses of 0, 25, 

125 and 250 mg/kg/day imazapyr showed no statistically significant effects on survival or other

endpoints monitored. The product registrant concluded the NOEL at 250 mg/kg/day, the HDT. 

Rats and mice fed imazapyr for 2 and 1.5 years (respectively) exhibited an increased incidence of 

congestion of the brain in females (mice), fluid accumulation in the air sacs of the lungs in females

(mice), increased incidence of kidney cysts in males (mice), increase in abnormal blood formation 

in the spleen (rats), increase of blood pooling in the liver (rats), increase in thyroid cysts (rats), and

a decrease in food efficiency (rats). The diets contained 0, 1,000, 5,000, or 10,000 ppm imazapyr. 

However, the results from these chronic dietary exposure studies revealed no significant

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differences amongst the treatment concentrations, and the EPA has concluded NOEL 

concentrations at the highest dose tested—in this case, 1,301 mg/kg/day in mice, and 503 

mg/kg/day in male rats (Fed Reg 62, 1997). Tumors were identified in both the high-dose and 

control groups at insignificantly different rates and the EPA therefore concluded the herbicide is

not considered to be carcinogenic. In vitro gene mutation studies using the Ames Salmonella

assay, chromosome aberration assay, point mutation assay, unscheduled DNA synthesis, and 

dominant-lethal assay yielded no significant results, and the herbicide was concluded to lack 

mutagenic activity (Fed Reg 62, 1997). 

Chronic concentrations of imazapyr at dose levels up to 1,000 mg/kg per day in rats and up to 400 

mg/kg per day in rabbits resulted in no significant differences amongst the doses tested for 

mutations or birth defects (Cyanamid Ltd. 1997). Once again, the NOEL for Arsenaltm in these 

developmental toxicity studies was reported at the HDT for rabbits (400 mg/kg/day), but lowered 

to 300 mg/kg/day for the rats, as the highest dose tested (1,000 mg/kg/day) increased salivation in 

the gravid females, even though no specific developmental toxicity endpoint was altered in the rats. 

No intergenerational effects were observed in a subsequent two-generation rat study conducted at

dietary concentrations up to 10,000 ppm (738 mg/kg/day) and the HDT was again accepted as the

NOEL. 

3.3.2 Birds 

The reported acute oral LD50 concentration for bobwhite quail and mallards is >5,000 mg/kg-diet. 

The reported subacute oral LD50 concentration for bobwhite quail and mallards reproduction is 

>1,890 mg/kg-diet. These values represent the highest doses tested to date. No significant 

bioaccumulation has been reported in avian species. Results from past avian studies conducted by the 

product registrant with Arsenaltm and/or technical grade imazapyr are summarized in Table 3-10. 

Table 3-10: Lethal and Sublethal Toxicity of Arsenal in Controlled Avian Experiments. 

Species/Test LD50 or LC50 NOEL (of test substance) 

Northern Bobwhite Quail LD50/18-week dietary 

exposure effects on egg production, hatchability, 

& hatchling survival) 

> 1,890 mg/kg 

(~200 mg/kg-quail) 

1,890 mg/kg 

(~200 mg/kg-quail = the HDT) 

Mallard Duck LD50/18-week dietary exposure

effects on egg production, hatchability, & 

hatchling survival 

>1,890 mg/kg-diet 

(~200 mg/kg-duck) 

1,890 mg/kg-diet

(~200 mg/kg-duck) 

Northern Bobwhite LD50/5 day acute dietary 

exposure effects on survival) 

>5,000 mg/kg-diet 

(~674mg/kg-quail) 

5,000 mg/kg-diet

(~674mg/kg-quail) 

Mallard Duck LD50/test 850.2200 (5 day acute

dietary exposure effects on survival) 

>5,000 mg/kg-diet 

(~1149 mg/kg-duck) 

5,000 mg/kg-diet

(~1149 mg/kg-duck) 

Willapa Bay, Padilla Bay, and other locations in the state where Spartina has colonized provide

substantial and significant waterfowl habitat (see Appendix B for review of ecology in these areas). 

As such, testing with the mallard duck provides a good surrogate for other waterfowl species that

use these areas and could potentially be exposed to imazapyr after a Spartina treatment. 

Shorebirds also use these estuarine habitats, and Willapa Bay supports the most significant stopover point along the west coast for migrating shorebirds. Shorebirds are greatly affected by the

colonization of Spartina, as it reduces available foraging habitat for these species. Indeed, 

monitoring studies have shown that shorebirds will not use Spartina meadows to feed in (Patten 

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and Stenvall 2002). Notwithstanding, the toxicological risks to shorebirds must be considered. 

Unfortunately, we found no data to address the potential toxicity of imazapyr to shorebirds. One

study, however, examined the penetration force required to penetrate mudflat sediment colonized 

with Spartina relative to an uncolonized mudflat, and a colonized mudflat treated with tilling or 

herbicide. In that study, an artificial Dunlin beak penetrated a similar (insignificantly different)

distance in the tilled Spartina meadow as in the bare mudflat. Herbicide treated meadows killed 

the Spartina, but did not soften the sediment, like the tilling. These results suggest that the habitat

risks from Spartina infestation are not necessarily alleviated by herbicide treatment alone. 

untreated

sprayed

spray+m ow

tilled

m udflat

beak

probing

density

/0.25m 2

100

80

60

40

20

0

-20

Figure 3-11: Penetration distance in artificial beak tests. 

(Source: Patten and Stenvall 2002.)

In addition to the toxicological data gap on shorebird and raptor species, no data were identified on 

preening exposure or inhalation exposure potentials associated with imazapyr. 

3.3.3 Insects 

The reported acute contact LD50 toxicity concentration for the honey bee is approximately > 0.1 

mg imazapyr/bee (Gagne et al. 1991). SERA (1999) estimated this dose to be greater than 1,000

mg/kg-bw, assuming 100% absorption of the applied dose, and an average body weight of 0.093 

g/bee [0.1 mg/0.000093 kg = 1075 mg/kg-bee, or ~ 1,000 mg/kg-bw]. Similar to the avian and 

mammalian studies previously addressed, there was no dose tested that resulted in acute or chronic

mortality, and the NOEL for the bee was taken to be the highest dose tested, or 100 ug/bee (1075 

mg/kg). Using the mammalian toxicity criteria adopted by the EPA (Table 3-9), imazapyr would

be considered practically non-toxic to the bee. 

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In terrestrial environments, herbicide treatment has been shown to increase the local abundance of

arthropods—likely as a response to the increased food supply for these detritivores from the dead 

and decaying vegetation. Arthropods serve as a substantial, high-energy food source for many bird 

species, and this relationship holds true both for terrestrial birds, as well as waterfowl and 

shorebirds during periods of migration (Cohen et al. 2000). A careful examination of the use of 

dead and decaying (post-treated) Spartina has not been conducted; however, it is conceivable that a 

similar relationship would be observed during the decay process. 

3.3.4 Reptiles and Amphibians 

Toxicity information of imazapyr with regards to reptiles and amphibians was not found through 

standard literature search engines. 

3.4 Toxicity to Aquatic Receptors 

A wide range of freshwater and marine fish and aquatic invertebrates occur in the surface waters of

Washington. The type of species found depends upon such factors as water temperature, salinity,

pH, and flow conditions. Because of the diversity of aquatic habitats throughout the state no single

species is representative of all habitats where fish and invertebrates could be exposed to imazapyr. 

Although Washington State supports several Endangered Species Act (ESA)-listed aquatic species, 

these species are often not those that are used for toxicity testing. Indeed, most testing that has

been conducted has focused on freshwater species, or marine species that are not generally found

in Washington State’s estuaries . Thus, although the central focus of this risk assessment considers 

the potential use and risks from imazapyr when applied in an estuarine environment, the following

discussion on imazapyr’s aquatic toxicity considers the total weight of evidence acquired from

aquatic species from various systems. 

In this section we examine the acute and chronic toxicity studies conducted on imazapyr in the

aquatic environment. Specifically, the acute and chronic toxicity and accumulation potential of 

imazapyr to fish, aquatic invertebrates, and non-target aquatic plants are discussed. Acute data are 

generated by the conduct of 2 to 4 day-long studies under controlled conditions. Table 3-11 

provides general risk assessment standards for fish and aquatic invertebrates based on acute

toxicity LC50 concentrations. This guide was used as a preliminary template to gauge risks 

associated with project actions that are broad in scope, such as the use of imazapyr by WSDA for

agricultural management. Chronic effects in aquatic organisms are usually based upon adverse

reproductive or growth effects, such as decreased hatching success, decreased survival of larvae, 

decreased growth of larvae or juveniles, and decreased reproductive capability. Chronic effects 

may occur from either acute or chronic exposures. In the majority of studies reviewed in this

section, little or no chronic toxicity data were available because earlier tier acute testing did not

indicate the need for further data development. 

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Table 3-11: Toxicity Classifications to Address Acute Risk to Aquatic Organisms from

Chemical Use. 

Fish or Aquatic Invertebrates Hazard Category Acute Concentration LC50 (mg/L) 

Very highly toxic <0.1 

Highly toxic 0.1-1 

Moderately toxic >1-10 

Slightly toxic >10-100 

Practically non-toxic >100 

3.4.1 Fish 

The reported acute toxicity LC50 concentration for rainbow trout, bluegill sunfish, and channel 

catfish is >100 mg/L based on product registrant studies with technical grade imazapyr using 

standard 96-hr exposure studies (Mangels and Ritter 2000). Tests were also conducted with the 

Atlantic silverside to address the potential toxicity of imazapyr to marine fish. In those tests the

highest concentration tested was 184 mg/L, which yielded no significant toxicity (mortality). As

with studies with terrestrial animals, the NOEC was taken as the HDT, or 100 mg/L for freshwater 

fish and 184 mg/L for marine fish. On this basis, imazapyr was considered practically non-toxic to

freshwater fish based on toxicity criteria outlined in Table 3-11. 

Imazapyr has not been thoroughly tested for chronic or sub-lethal effects with a wide variety of

aquatic organisms, but those few tests conducted are worth summarizing. Early life stage survival

tests with rainbow trout and fathead minnow embryos and sac-fry continuously exposed to 

imazapyr revealed no effects on hatching or survival at concentrations as high as 92.4 a.i. mg/L

and 118 mg a.i/L, respectively. Again, these were the highest concentrations tested. A full life

cycle test with fathead minnow with concentrations up to 120 mg a.i./L also did not elicit toxicity. 

It is unclear why the product registrant did not pursue testing with higher concentrations to 

establish the true maximum tolerated dose. Such testing has applications when addressing 

potential spill scenarios with the highly soluble herbicide. However, recent results by University

of Washington researchers help to eliminate this uncertainty (C. Grue personal communication, 

2003). Grue and others examined the toxicity of imazapyr in 96-hr tank tests with juvenile rainbow 

trout (Table 3-12). As demonstrated in these tests, the concentrations required to achieve 50%

mortality are exceedingly high. Indeed, the concentration of the formulations required exceed the

total salt concentration of full strength sea-water (typically 30,000 to 38,000 mg/L). The NOEC 

concentrations have not been calculated from this work as of the publication date of this 

assessment. As one purpose of the studies was to compare the toxicity of imazapyr with Rodeotm

(glyphosate), data are summarized for this herbicide as well. As further demonstrated in Table 3-

12, the LC50 of glyphosate established in the same trials was approximately two orders of 

magnitude more toxic than the Arsenaltm herbicide. 

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Table 3-12: 96-hour LC50 Values with 0.3 g juvenile rainbow trout exposed to imazapyr

(Arsenal) or glyphosate (Rodeo) tank mixes. 

(Source: C. Grue 2003, personal communication.) 

Product Tested LC50 of Concentrate LC50 Expressed as Active Ingredient 

Arsenal Herbicide 77,716 ppm Arsenal 

(72,183-72,243)*

22,305 mg/L imazapyr 

(20,718-20,891)*

Arsenal Concentrate 43,947 mg/L (41,446-46,408)* 23,336 (22,024-22,643)* 

Rodeo 782 mg/L (719-845)* 782 (719-845)* 

* 95% confidence interval of four replicated trials with geometrically arranged concentrations and a negative control.

Sub-lethal endpoints other than the early-life-stage and life cycle tests conducted with the standard

test species have not been fully explored with imazapyr. One recent study examined the potential 

for imazapyr (Arsenal) and glyphosate (Rodeo) to elicit micronuclei in the African cichlid fish 

(Tilapia rendalli) abdominally injected with the herbicides (Grisolia 2002). Micronuclei have been 

proposed as a reliable indicator of environmental mutagenesis in aquatic and terrestrial animals,

and have been evaluated in a variety of mollusc, fish and amphibians as an indicator of potential

mutagenicity (Al-Sabti and Metcalfe 1995, Vernier et al. 1997). Micronuclei are reflected as 

chromosomal abnormalities in blood smears. However, the significance of elevated micronuclei

frequency at the population level has not been fully determined. In the Grisolia (2002) study,

significantly elevated numbers of micronuceil were observed following imazapyr exposure, but

only at 80 mg/kg-bw, the maximum tolerated dose (MTD). Evidence of sub-lethal effects at the 

MTD are not considered valid indicators of sub-lethal toxicity, as the fish are exhibiting overt

cytotoxicity (cell death) signs. Chromosomal aberrations such as micronuclei are common during 

cell death; their significance to mutagenicity studies is relevant when occurring as a sub-lethal

toxicological response to chemical exposure doses below those which cause cell death. 

Threatened and Endangered Fish Species Hazards 

Resident fish populations managed by the USFWS under the Endangered Species Act (ESA) are

delineated as “distinct population segments” (DPS), while the NOAA-Fisheries, which manages

marine and anadromous ESA-listed stocks, delineates populations as “evolutionarily significant 

units” (ESUs). Addressing the uncertainty posed by using surrogate test species that may or may not

be as sensitive as the threatened and endangered (T&E) populations unique to an area is always 

problematic in ecological risk assessments. For example, Mayer and Ellersieck (1986), in their 

compilation of an acute toxicity database for 410 chemicals tested on aquatic organisms, found that

toxicity amongst species could range by as much as five orders of magnitude, and for a given 

species, toxicity could range by as much as 9 orders of magnitude. This data base, however, lacked

critical review to thoroughly filter test comparisons that were appropriate (e.g., standardized test

conditions). In Willapa Bay, where the Spartina infestation is the greatest, no fish populations are

considered “threatened” or “endangered” under the ESA. In Puget Sound locations where Spartina

has colonized, the native chinook salmon and coastal bull trout are both considered threatened, with

the former managed by the NOAA-fisheries, and latter by the USFWS. 

No imazapyr toxicity tests have been conducted with DPS or ESUs’ of fish listed as threatened or 

endangered in the State of Washington. Although testing has not been conducted with these specific

native populations that may utilize Spartina infested habitats in Washington, the standard testing 

conducted with the closely related rainbow trout discussed in the preceding section provides a good 

surrogate for predicting survival effects in these closely related native stocks. Toxicity testing under 

FIFRA requires data to be collected on surrogate species that addresses acute toxicity, embryo-larval 

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survival, and life-cycle tests. As indicated by those tests, imazapyr would be considered practically 

non-toxic on the basis of results in rainbow trout, bluegill and fathead minnow. 

Studies conducted in the early 1970s examined the sensitivity of four fish families to 65 different 

chemicals (Macek and McAllister 1970); salmonids were the most sensitive of the four families 

(12 species) represented. A more recent study by Sappington et al. (2000) evaluated the 

comparative sensitivity of eight ESA-listed fish species to standard test organisms exposed to 

five different pesticides or metals in order to validate the use of surrogate species as a predictive

tool in toxcilogical assessments. Acute 96-hr exposure trials were conducted, but none of the 

chemicals tested by these authors were herbicides, and all but nonylphenol had had significant 

previous testing. The sensitivity of listed cold-water species tested (Apache trout, Lahontan 

cutthroat trout, greenback cutthroat trout) did not differ significantly after 96-hr exposures from 

rainbow trout for copper, nonylphenol, or carbaryl. However, they were significantly more

sensitive to the organophosphate permethrin and pentachlorophenol than the rainbow trout. 

Toxicities exhibited throughout the testing varied with chemical, with some listed species 

exhibiting greater or lesser sensitivity than the standard test species at some time points (e.g., 12 

hours). Although differences were documented which were sometimes statistically significant 

depending on the time point, the listed species were not always the most sensitive. Most 

importantly, the maximum degree of difference recorded was less than two-fold, except

pentachlorphenol and permethrin for which the listed species exhibited LC50 concentrations less 

than half of the surrogate rainbow trout. The authors concluded that a safety factor of two would 

provide a conservative estimate in risk assessments for listed cold-water, warm-water and 

euryhaline fish species based on these findings. 

Another common criticism of ecological risk assessments relying on surrogate species to address

potential T&E species effects is the lack of data on sublethal endpoints of site-specific relevance. 

The coastal estuaries where imazapyr could be applied to control Spartina serve as a primary staging

area for salmon smolts that are migrating to the sea to mature. The brackish salinties found in the 

estuaries provide a range of salinities salmon smolts use to adapt to full strength sea-water. The 

osmoregulatory capacity has been used as one test to establish whether a chemical might affect this

sensitive life stage. Patten (2003) examined this capacity, measured as plasma sodium level and gill 

ATPase activity in a 24-hr seawater challenge, in chinook salmon smolts exposed to imazapyr 

concentrations up to 1.6 mg/L (Figure 3-12). This maximum test concentration was over 470-fold 

greater than the maximum water concentration recovered in the companion study where imazapyr 

was applied to bare-mud and measured in waters from the first tidal wash (Patten 2003). As

demonstrated in Figure 3-12, there was no consistent dose-response effect recorded on these 

endpoints of sublethal physiological relevance. 

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Figure 3-12: Plasma sodium and gill ATPase activity of chinook salmon exposed 

to imazapyr. 

(source: Patten 2003.)

3.4.2 Aquatic Invertebrates 

One study where Arsenal was applied with a surfactant (not defined) with the freshwater water flea

(Daphnia magna) yielded a 48-hr LC50 of 350 mg-Arsenal/L (79.1 mg ae imazapyr/L) and an

NOEC of 180 mg-Arsenal/L (40.7 mg ae/L). Other product registrant studies where Daphnia was 

exposed to an imazapyr formulation (~50%) lacking the surfactant produced a 48-hour EC50

concentration of 373 mg a.e./L (Cyanamid 1997). The results of these two studies highlight the

potential effect of surfactant on aquatic toxicity, and the authors concluded that the, “components of

the Arsenal formulation, other than a surfactant, do not influence the toxicity of imazapyr to aquatic

organisms.” Kintner and Forbis (1983) also reported 24 and 48-hour LC50 concentrations of greater 

than 100 mg/L (the HDT), in static tests conducted with newly-hatched Daphnia (less than 24 hours 

old). Chronic studies have also been conducted with the water flea (Manning 1989). In that study, 

no adverse effects on survival, reproduction or growth of 1st generation Daphia were recorded after 

7, 14 and 21-days of exposure at concentrations up to 97.1 mg/L, the HDT. Per FIFRA registration 

requirements, the NOEC was considered to be the HDT (97.1 mg/L), and the maximum allowable

toxicant concentration (MATC) was considered to be > 97.1 mg/L. 

Testing with other invertebrate species that exhibit alternative life cycles has been limited to 

growth studies with the Eastern oyster (Crassostrea virginica), and survival of pink shrimp.

Although these species are not native to coastal Washington, they do provide reasonable surrogates

for the Pacific oyster (Crassostrea gigas) and burrowing shrimp (Neotrypaea sp.) that are common

to areas where Spartina has become established. In these product registrant tests, the EC50 for

growth inhibition was established at a concentration greater than 132 mg-imazapyr/L, with the

NOEC set at this concentration—the HDT. The pink shrimp survival LC50 was > 189 mgimazapyr/L, and the NOEC was again set at this HDT (Mangels and Ritter 2000). 

3.4.3 Non-target Aquatic Vegetation 

Native salt marsh plants and algae resident to the estuarine environments where imazapyr could be

applied have the potential to be negatively affected by the broad spectrum herbicide, and a range of 

studies by both the product registrant and others document this possibility. Table 3-13 summarizes

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product registrant studies with a variety of freshwater and marine algae and aquatic plants exposed

to either technical grade imazapyr, or to Arsenal with surfactant. As indicated in Table 3-13, 

toxicity to the vascular plant duckweed was nearly three-orders of magnitude greater than the

toxicity associated with the algal species tested. Notably, the toxicity of the Arsenal formulation 

did not differ from that of the technical grade imazapyr for duckweed, although it was found to be 

approximately 5-fold more toxic to green algae. 

Table 3-13: Toxicity of Technical Grade Imazapyr and Arsenal* (with surfactant) to Algae

and Aquatic Plants, as Established Through Controlled Product Registrant Studies. 

(Source: Mangells and Ritter 2000.) 

Species/Test EC50 EC25

Green Algae Growth (Selenastrum capricornutum) 71 mg/L

14.1 mg/L* 

48 mg/L 

8.36 mg/L* 

Freshwater diatom (Navicula pelliculosa) > 59 mg/L > 59 mg/L

Saltwater diatom (Skeletonema costatum) 85.5 mg/L 42.2 mg/L 

Blue-green algae (Anabaena flos-aquae) 11.7 mg/L 7.3 mg/L

Duckweed (Lemna gibba) 0.024 mg/L

0.0216 mg/L* 

0.013 mg/L 

0.0132 mg/L* 

Recent studies conducted by Patten (2003) also document the potential for imazapyr to impact

non-target vegetation in those areas where Spartina control is envisioned. In this study, the effects 

of imazapyr were examined on the non-native Japanese eelgrass (Zostera japonica) and compared

against glyphosate. This species of eelgrass dominates the intertidal zone unlike the native eelgrass

(Z. marina) which is primarily found sub-tidal. For both herbicides the eelgrass canopy was killed 

if herbicide was applied on dry eelgrass at low tide, although the imazapyr was more toxic. If

applied with a film of water overlying the bed, then no effect was recorded. Within 12 months

post-treatment, all impacted eelgrass beds had recovered. There was no difference in toxicity over

the range of doses tested, (0.84 kg ae/ha and 1.68 kg ae/ha). Persistence was not recorded in the 

sediment underlying these eelgrass beds, hence resistance to the establishment of native salt marsh 

plants such as Salicornia was not considered a risk. 

3.5 Adjuvant and Inert Ingredient Toxicity to Terrestrial and Aquatic Ecological 

Receptors

Adjuvants are carriers mixed with herbicides that increase the binding and/or uptake of the

herbicide into target plants. Typical adjuvants include surfactants and crop oils that are mixed with 

the herbicide prior to application. Inert ingredients are components within the patented herbicide

product formulations that are reported to have no herbicidal activity. Current FIFRA regulations do 

not require manufacturers to reveal the surfactant formulations, as FIFRA regulates the active

ingredients only. Similarly, many of the inert ingredients in the commercial formulations of the 

various imazapyr products on the market are not known. Herbicide toxicity studies conducted 

under FIFRA are required to evaluate the active ingredient of the product formulation only, and not 

the toxicity of the “inert ingredients” or the surfactants that may be used to facilitate plant 

adsorption and uptake of the herbicide. For some ecological receptors, particularly aquatic

receptors, the choice of which surfactant is used to administer the herbicide can have substantial

ecological relevance, as the few tests conducted with surfactants have shown higher toxicity than 

the herbicide. Similarly, in environments where a variety of herbicides and/or pesticides may be

used, the potential for chemical interactions of inert ingredients should also be understood to 

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minimize risks. This section of the hazard assessment therefore attempts to summarize the existing 

information on the toxicity inherent to the inert ingredients and surfactants that could be used in the 

application of imazapyr to control Spartina. 

3.5.1 Inert Ingredients 

Two of the inert ingredients in Arsenal® are listed as glacial acetic acid (CAS #64-19-7) and water 

(CAS #7732-18-5) (NCAP 2003). Water is non-toxic and required for life. The toxicity of acetic

acid is tabulated below (Table 3-14), as summarized by Merck (1989) and Verchueren (1983). 

Acetic acid is also a component of LI 700, a common non-ionic surfactant with potential use with

imazapyr. 

Table 3-14: Acetic Acid Toxicity to Ecological Receptors. 

Test species 

Class of 

organism Toxicity test Toxicity end point Value Unit 

Wheat Plant EC 50 Visible injury 23.3 mg/m3

Alfalfa Plant EC 50 Visible injury 7.8 mg/ m3

Corn Plant EC 50 Visible injury 50.1 mg/ m3

Pseudomonas 

putida 

Bacteria Toxicity threshold Multiplication inhibition 2850 mg/l

Microcystis 

aeruginosa 

Algae Toxicity threshold Multiplication inhibition 90 mg/l

Scenedesmus

quadricauda 

Green algae Toxicity threshold Multiplication inhibition 4000 mg/l

Entosiphon sulcatum Protozoa Toxicity threshold Multiplication inhibition 78 mg/l

Uronema parduczi Protozoa Toxicity threshold Multiplication inhibition 1350 mg/l

Vorticella campanula Protozoa Toxicity threshold Perturbation level 12 mg/l

Brine shrimp Arthropoda TLm* 32-47 mg/l

Grammarus pulex Arthropoda TLm* 6 mg/l

Limnea ovata Mollusca Perturbation level 15 mg/l

Bluegill Fish TLm* (24, 96-hr--

respectively)

100-1000,

75 

mg/l

Mosquito fish Fish TLm (24-96 hr) 251 mg/l

Fathead minnow Fish LC50 (1, 24, 48, 72, 

96-hr--respectively) 

Death 175, 106,

106, 79, 79) 

mg/l

Culex sp. larvae Insects TLm (24-48 hr) 1500 mg/l

Mice Mammals LC50 (l hr) Inhalation 5000 ppm 

*median tolerance limit 

3.5.2 Surfactants

Surfactants are used to reduce the surface tension of water, enabling a “bridge” to form between 

two chemicals or media that would not normally mix (e.g., oil and water). When used with 

herbicides, they are intended to maximize the amount of spray solution that sticks to the leaf

surface, and hence increase uptake. Surfactants commonly used to promote imazapyr and 

glyphosate adsorption and uptake are generally of two classes: non-ionic nonylphenol alcohols

and/or fatty acids, and crop-oil based concentrates. Studies evaluating the efficacy of imazapyr and

glyphosate with various surfactants have revealed few differences in the efficacy of the herbicides 

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based on the surfactant (Patten 2002). All surfactants tested with imazapyr provided effective 

control, but R-11, the approved surfactant for use with glyphosate, was not tested with imazapyr, 

making a direct comparison difficult (Table 3-15). However, the author states that “application

made with short dry time might better distinguish surfactant effects than did these trials, all of 

which had ample dry time”. 

Table 3-15: Effect of surfactant applied in September 1999 and 2000 on the efficacy of

imazapyr for smooth cordgrass control in Willapa Bay, WA. 

(Source: Patten, 2002.)

Percent control 13 months after treatment 

Herbicide 

Rate 

(kg/ha) Surfactant 

Percent 

(v/v) Site 1 Site 2 Site 3 

Imazapyr 1.68 Agri-Dex 1.0 99 85 96

1.68 Agri-Dex 2.0 96

1.68 Hasten 1.0 100 83 94

1.68 Kinetic 0.5 89

1.68 Dyn-Amic 1.0 96

1.68 Syl-Tac 1.0 92

Glyphosate 8.4 R11 1.0 69 85

Untreated na na Na 0 0 0

 

Although there appears to be little difference amongst surfactants in their potentiation of herbicide

efficacy, their inherent chemical properties can have a range of environmental issues that are 

independent of the herbicide formulation they may be applied with. For this reason, it is prudent to 

examine their properties and toxicity independently. Table 3-16 summarizes descriptions of 

surfactant environmental fate, chemistry and toxicity as provided in the original EIS and obtained 

from the manufacturer’s material safety data sheets (WSDA 1993E, chapter 11.00). In brief, the

acute toxicity of alkylphenol ethoxylate surfactants like R-11tm and X-77 tm to fish and other 

aquatic species has been reported in the range of 4 to 12 mg/L. Acidifying agents like LI-700, and 

crop-oil based surfactants like Hasten tm and Agri-Dex tm exhibit lower toxicity. On the basis of

EPA aquatic toxicity criteria, all the surfactants used would be considered practically non-toxic

(LI700 tm, Hasten tm and Agri-Dex tm ) to moderately toxic (R-11, X-77). All of the surfactants can 

cause irritation to skin and ocular tissue at high doses, and receive ratings of moderate (scores of 4 

to 6 on an 8 pt scale) irritation in mammals (Table 3-17). By oral administration, the limited 

testing done with the surfactants in mammals indicates they would classify as “practically nontoxic”. 

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Table 3-16: Chemistry and Fate of Surfactants Potentially Used With Imazapyr and 

Glyphosate.

Surfactant 

Known Ingredients* & 

Surfactant Class Chemical Properties 

Degradation Rate and 

Pathway 

General Toxicity 

Rating* 

R-11tm (surface

activator), Wilbur-Ellis 

Co. 

Isopropyl (butyl) alcohol

20%, nonionic surfactants 

80% (octyl phenoxy 

polyethoxy), silicone. 

Class: Nonionic 

alkylphenol ethoxylate 

Soluble in lipid & water, 

Flammable,

Spec. Gravity = 1.0 

Slowly biodegraded by

progressive shortening

of ethoxylate chain; 

intermediate breakdown 

products of polytheylene

glycol (anti-freeze) and 

short-chain ethoxylates.

Mammals: practically

non-toxic orally, mild skin 

irritation possible 

Fish and other aquatic 

biota: moderately toxic 

LI-700 tm (penetrating

surfactant), Loveland

Industries, Inc.

Phosphatidylcholine

(lecithin) at 800 g/L,

propionic acid, and

alkylphenyl 

hydroxypolyoxyethylene

Class: Acidifying agent 

Soluble in lipid & water, 

Not Flammable 

Spec. Gravity = 1.03 

Biodegradation

presumed rapid due to

natural lecithin

ingredients.

Mammals: practically

non-toxic orally, but 

causes skin irritation 

Fish and other aquatic 

biota: moderately toxic 

X-77 tm (spreader 

activator), Valent

Corp. 

Alkylarylpoly 

(oxyethylene), glycols, free

fatty acids, isopropyl

alcohol. 

Class: Nonionic 

alkylphenol ethoxylate

Soluble in lipid &water, 

Flammable

Slowly biodegraded by

progressive shortening

of ethoxylate chain; 

intermediate breakdown 

products of polytheylene

glycol (anti-freeze) and 

short-chain ethoxylates.

Mammals: practically

non-toxic orally

Fish and other aquatic 

biota: moderately toxic 

HASTEN tm Proprietary: fatty acids 

from seed oils esterified

with alcohol 

Class: oil based surfactant 

Non-ionic, dispersible in

water as micelles, but 

unknown solubility. 

Sp. Gravity = 0.9 

Biodegradation

presumed rapid, but no

formal studies conducted

of which we are aware. 

Mammals: practically

non-toxic through oral 

routes 

Fish and other aquatic 

biota: slightly toxic 

AGRI-DEX tm Proprietary: heavy range

paraffin-based petroleum 

oil with polyol fatty acid

esters and 

polyethoxylyated 

derivatives

Class: oil based surfactant 

Dispersible in water

(forms micelles), 

moderate flammability, 

Biodegradation

presumed rapid, but no

formal studies conducted

of which we are aware 

Mammals: practically

non-toxic through oral 

ingestion, mild skin and 

eye irritant, 

Fish and other aquatic 

biota: practically nontoxic 

*See tables 3-9 and 3-12 for toxicity classification schemes 

Past studies with glyphosate have shown that the toxicity of surfactants is generally greater than 

the toxicity of the herbicide formulation or active ingredient alone. For example, studies with

Rodeo formerly discussed in the original EIS relate how the toxicity of the Rodeo formulation was

1,100 mg/L without surfactant, and 680 mg/L with the mixture containing 0.4 percent X-77 

(Mitchel et al 1987). A similar relationship has been observed with aquatic invertebrates with

Rodeo (Henry 1992). Recent studies with both imazapyr (Arsenal) and glyphosate (Rodeo) 

examined the inherent toxicity of the surfactants also, both with and without the herbicides (Smith 

et al. 2002, unpublished data). As demonstrated in Table 3-17, the toxicity of the seed and crop-oil 

based surfactants Hasten and Agri-Dex to rainbow trout was two to three orders of magnitude

lower (respectively) than R-11 in this study. When surfactant was mixed with herbicide, the 

toxicity of the surfactant was reduced and the toxicity of the herbicide was increased. These

studies reveal that the toxicity associated with herbicide/surfactant mixtures is not additive, and is 

generally associated with the surfactant. Of the surfactants examined in detail, the order of

toxicity, from lowest to highest, would appear to be as follows: Agri-Dex, Hasten, LI700, X-77

and R-11. It is noteworthy, that only R-11, the surfactant that appears most toxic from the recent

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tests, is approved for use with glyphosate in the estuarine environment where herbicide treatment 

of Spartina is conducted. 

Table 3-17 Toxicity of Surfactants With and Without Herbicide. 

(Sources: Smith et al. 2002, Henry 1992, Mitchell et al. 1987, WSDA 1993E.)

Chemical Tested Mammalian Toxicity LD50 (ppm) Aquatic Toxicity (ppm) 

R-11 surfactant 5,840 oral, 13000 dermal (rabbit) 6.0, rainbow trout 96-hr LC50* 

4.2, bluegill sunfish 96-hr LC50

LI-700 surfactant >5,000 oral, 5,000 dermal (rat) 17, rainbow trout 96-hr LC50* 

22, rainbow trout 24-hr LC50* 

210, bluegill sunfish 96-hr LC50 

190, daphnia 48-hr LC50 

Hasten surfactant No Data 74, rainbow trout 96-hr LC50* 

98, rainbow trout 24-hr LC50*

Agri-Dex surfactant >5,010 oral (rat), > 2,020 dermal (rabbit) 271, rainbow trout 96-hr LC50* 

386, rainbow trout 24-hr LC50*

X-77 surfactant > 5,000 oral (rat), > 5,000 dermal (rabbit) 4.2, rainbow trout 96-hr LC50 

4.3, bluegill sunfish 96-hr LC50 

2, water flea (daphnia) 48-hr LC50

Rodeo (as glyphosate) 3,800 oral, 5,000 dermal (rabbit) 580, rainbow trout 96-hr LC50 

545, water flea (daphnia) 48-hr LC50

Rodeo + X-77 No Data 130, rainbow trout 96-hr LC50 

130, water flea (daphnia) 48-hr LC50

Rodeo + R-11 No Data 5.4, mg/L rainbow trout 96-hr LC50*

Rodeo + LI700 No Data 23, mg/L rainbow trout 96-hr LC50*

Arsenal + Hasten No Data 113, mg/L rainbow trout 96-hr LC50

Arsenal + Agri-dex No Data 479, mg/L rainbow trout 96-hr LC50*

*Unpublished data from Smith et al. (submitted to Bull. Env. Of Contam. And Tox.). Data represents mean of 4 trials, upper 95%

confidence limit within 5 to 20% of mean over all herbicide trials (not shown) 

The non-ionic alkylphenol derived surfactants may pose additional hazards beyond the evidence 

provided in acute toxicity tests. The alkylphenols and octyl phenol ethoxylates belong to a broader 

class of chemicals known as the “nonylphenols”. It has been estimated that approximately 80 

percent of the alkyl phenol ethoxylates are nonyl phenol ethoxylates and the other 20 percent are

octyl phenol ethoxylates (Cox 1998). Because these compounds are not part of the herbicide

formulation, their exact formulations are patent protected and are not reportable under FIFRA. 

However, the EPA considers the nonylphenols as an “inert of toxicological concern.”

Nonylphenol ethoxylates degrade to nonyl phenol and related compounds that can be somewhat

persistent in the environment. Sublethal effects at exposure concentrations below acutely toxic

level were previously described (WSDA 1993E) and included impaired swimming activity, altered 

breathing rate, and reduced heart rate in fish at 0.5 mg/L, and inhibited siphon retraction, byssal

thread formation and reduced burrowing activity in sessile shellfish at concentrations greater than 1

mg/L. Lethal effects as reported in the literature are summarized in Table 3-18. The intermediate

breakdown products of these surfactants can include both linear and branched chain alkylphenols, 

which may also have inherent toxicity. Some of these products have been shown to elicit weak

estrogenic effects when administered at high doses to laboratory animals (reference). Determining 

the actual quantity of alkylphenols in each surfactant formulation, and their potential

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environmental concentrations and risks is not entirely possible because the proportions in each 

surfactant formulation are not known. 

Table 3-18: Acute toxicity of nonylphenol to aquatic biota.

Test species 

Class of 

organism Toxicity test Toxicity end point Value Units 

Mytilus edulis1 Mussel Bioconcentration

Factor 

NA 10 Wet weight

Caenorhabditis 

elegons2

Nematode LC50 (24 hr) Death 7.2 mg/l

Mysidopsis bahia2 Mysid LC50 (96 HR) Death 43 mg/l

Fathead minnow2 Fish LC50 (96 HR) Death 135 mg/l

Gadus morhua2 Fish LC50 (96 HR) Death 3000 mg/l

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October 2003 Page 56
4.0 EXPOSURE ASSESSMENT 

In this chapter we characterize the potential exposure to fish and wildlife receptors from the use of 

imazapyr to control Spartina based on exposure parameters generated from current and projected 

application practices. Wildlife species are predominantly exposed to herbicides by consuming 

treated vegetation and/or water, and/or by transfer of the chemical through natural food chains. 

Contact (dermal) and inhalation exposure can provide additional, although marginal, exposure.

Inhalation exposure of drift is generally extremely limited because application equipment creates

noise that causes mobile birds and mammals to avoid the immediate area. Nocturnal animals such

as the rat would largely avoid inhalation exposure because WSDA applies herbicides only during 

daylight hours. For this assessment drinking water exposure would also be limited because

treatments will occur in estuarine waters where freshwater is limited, although rain is generally 

abundant, and consumption of freshwater accumulated on the plants is possible. Our ability to

characterize imazapyr exposure for some sectors of wildlife such as amphibians and reptiles is 

limited because of a lack of basic biological (life history) and toxicological information. Site

specific studies would be required to address quantitative risk assessment in each area where 

Spartina has infested, so exposure modeling was conducted in lieu of site specific work to gauge

exposure doses based on application rates and delivery mechanisms of relevance to wildlife

receptors. Exposure doses were then compared to reference doses from the toxicity literature to 

gauge the potential toxicity to relevant ecological receptors. 

4.1 Estimated Environmental Exposure Concentrations (EEC) 

The exposure concentrations or dose experienced by biota will differ by media (i.e., air, water,

food, and sediment), the habitat they use, how frequent they use the habitat, and the application 

rates of impazapyr. Estimates of environmental exposure concentrations (EEC) of imazapyr can 

be derived from empirical studies, and also from modeling, although the former is preferred. This

section summarizes the exposure concentrations that have been measured empirically, and those

from the literature that have been developed through modeling exercises. Table 4-1 summarizes 

empirical results where imazapyr was detected in environmental media. The derivation of many of 

these values was summarized in Chapter 3 of this document. The discussion below provides

assumptions applicable to EEC and exposure modeling. 

4.1.1 Application Rate 

For Spartina control, only the Arsenaltm imazapyr formulation is projected for use, however, no 

product endorsement is assumed. To model the estimated environmental concentrations of 

imazapyr in Arsenaltm, the different methods used for application must be understood. This

includes application rates, frequency, application volumes, and interception rates. The following 

assumptions were used regarding application rates. 

• Although lower rates may be used in certain areas, for ecological exposure and risk 

interpretations we assume that Arsenal will be applied at the maximum concentration 

recommended on the manufacturers label for aquatic use—6 pints Arsenal/acre. Six pints

per acre is equivalent to 1.5 lbs active ingredient (acid equivalents)/acre = 0.68 kg/acre = 

1.68 kg/ha. 

• Applications will occur a maximum of one time per year until eradication is complete. 

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October 2003 Page 57
• The neat herbicide formulation will be diluted with water and surfactant prior to 

application. Surfactant will be added to the herbicide/water mixture to yield 1% of the 

spray solution applied. 

• Three methods of herbicide administration are possible for Spartina control: (1) hand-held 

sprayer unit, (2) boom-mounted sprayer, and (3) aerial sprayer. Spray volumes by these 

methods can vary from a minimum of approximately 2.5 gallons/acre to a maximum of 80 

gallons/acre. 

• Herbicide quantity (mass) per unit area will not vary by spray volume (i.e., 1.5 lb/acre) but 

surfactant rates will, as these are normalized to spray volume. Ultra-low to low spray 

volumes of 2.5 to 20 gallons/acre are the most likely application rates, but risks of 

surfactant toxicity are also considered with high volume applications up to 80 gallons per 

acre. 

4.1.2 Water Concentrations 

Concentrations that reach the water will be affected by Spartina canopy interception, adsorption

onto the Spartina , uptake into the root zone, and aerial drift. However, the “worst case” imazapyr

water concentrations are assumed by considering no adsorption to sediment and/or vegetation, no 

foliar interception, and complete solubility on an incoming tide. Under these highly conservative 

conditions, water concentrations will change with depth, as represented in Figure 4-2. As

demonstrated in this figure, the modeled water concentration at the lowest depths evaluated are

consistent with an incoming tidal prism and are also reflective of the empirical results of Patten 

(2003) discussed earlier and summarized in Table 4-1. Under the worst case scenario, where the 

herbicide was applied to bare-mud, Patten (2003) projected extremely rapid dissipation from the

equation f = 0.0015 exp((319/x+38)). These field experiments suggested that applied imazapyr

would not be measurable after approximately 40 hours. 

Under typical treatment conditions the Spartina canopy will intercept herbicide, and will thus

effectively titrate the herbicide into the incoming waters as they rise over the Spartina canopy. The

highest concentration of applied herbicide will be deposited in the upper canopy and hence will not

be solubilized until the water depth reaches this portion of the canopy, allowing for greater dilution 

than would be expected if the herbicide was distributed uniformly on the plants. Although the 

herbicide is highly soluble, adsorption and uptake into plant matter is facilitated by surfactant and 

will also reduce the available herbicide for solubilization under typical applications. 

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October 2003 Page 58
Figure 4-1: Mud-boat applicator used to apply herbicide in Willapa Bay to control 

Spartina.

Linders at al. (2000) proposed a foliar interception rate of 40% in grasses based on a recent review

of retention values from pesticide and herbicide applications throughout the world. Patten (2003)

has proposed a foliar interception rate of 75% for Spartina treated with imazapyr based on 

empirical results, and this same value has been proposed by the manufacturers of Arsenaltm

(Mangels et al. 2000). High interception rates will maximize potential exposure for terrestrial 

herbivores, and minimize potential exposure to aquatic receptors whereas the reverse is true when 

interception rates are lower. 

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October 2003 Page 59
0

1

2

3

4

5

6

7

8

0 0.2 0.4 0.6 0.8 1

Water Depth (m)

Est. Water Concentration 

Imazapyr (mg/L)

Figure 4-2: Estimated water concentrations of imazapyr in tidal waters with no 

canopy interception and an application rate of 1.5 lbs/acre (0.68 kg/acre). 

4.1.3 Expected Plant and Animal Residues 

As discussed above, interception rates will affect both plant residues as well as potential water

concentrations. For Spartina clones, the Linder et al. interception value of 40% is more realistic 

with field applications because of the greater amount of edge around the clones and potential for

greater drift off the application site. For Spartina meadows, higher interception rates are more

likely, and the 75% interception value of Patten (2003) will be considered valid. Where the foliar

interception rate is 40%, it is assumed that 10% of the non-intercepted imazapyr will drift off-site

(or on non-target vegetation) and the remaining 50% will make contact with underlying sediment

and be solubilized upon the first tidal wash. These latter values were derived from studies

conducted in grasslands in the Netherlands during the growing phase (USES 2.0, 1998). 

Pesticide residues following applications of 1 lb/acre were estimated in a variety of plants and 

insects by Hoerger & Kenaga (1972) and more recently recorded from empirical data by Fletcher

et al. (1984). In the Hoerger and Kenaga study, tall grass had estimated average residue

concentrations of 73 mg/kg and estimated maximum concentrations of 87 mg/kg. In the 1984 study

by Fletcher and others, tall grass had average empirical residues of 29 mg/kg, and estimated 

maximum concentrations of 87 mg/kg. At the 1.5 lb/acre rate proposed for Spartina control with

imazapyr, the estimated average residue concentration detectable shortly after spraying would be 

approximately 110 mg/kg based on the model of Hoerger and Kenaga, and 43 mg/kg based on the

empirical results of Fletcher et al. (1984) (extrapolated to the higher application rate). The 

estimated maximum residue would be 131 mg/kg under both studies (Table 4-1). No field data on 

imazapyr residues in treated Spartina meadows or clones were available for review to compare 

against these residue estimates. 

The residue of imazapyr in plant tissues will change over time, and this degradation has not been 

examined empirically in treated Spartina. For the chronic dietary exposure assessments described 

in Section 4.2, the concentration in plant tissues over time was therefore modeled for this portion

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of the assessment using the methods of SERA (2001). Empirical residue concentrations in

Spartina over time following treatment remain a source of uncertainty. 

Table 4-1: Summary of Maximum and Average Imazapyr Detections in Relevant 

Environmental Media with Application Rates of 1.5 lbs a.i./acre.

Environmental Media 

Maximum 

Measured 

Average Maximum Measured, 

or Estimated Maximum* 

Average of all samples 

Measured, or Typical

Estimated*

Surface Water 5.77 mg/L[1a] 3.40 mg/L[1b] 0.1 mg/L[1c]

Pore Water 3.29 mg/L[2a] 2.94 mg/L[2b] 0.042 mg/L[2c]

Treated Sediment with no 

Overlying Canopy

5.7 mg/kg[3a] 5.4 mg/kg[3b] 3.2 mg/kg[3c]

Sediment Under Treated

Canopy 

2.27 mg/kg[4a] 1.42 mg/kg[4b] 1.4 mg/kg[4c]

Plant Tissue (long grass) No data 131 mg/kg[5a] 43 mg/kg[5b], 110 mg/kg[5c]

Animal Tissue No data No data < 50 ug/kg-wet wt. 

[1] a. First tidal flush, 0.6m outside immediate spray zone atop bare mud with no foliar interception, 3.5 hrs post trtmnt; b—average of 3 samples taken at first 

flush 0.6 m outside spray zone; c—geometric mean of all samples collected 3.5 to 4 hours post treatment at 0.6, 6.0 and 60 m outside spray zone (Patten 2003) 

[2] a—Highest value of three replicates taken 1 hr post treatment; b—average of three samples; c—average of three samples taken in sediment under canopy 15

days after treatment. (Patten 2003) 

[3] a—1-hr post treatment within treatment zone of bare mud; b—1-hr post treatment, average of 3 samples; c—geometric mean of all samples collected 3.5 to 4

hours post treatment at 0.6, 6.0 and 60 m outside spray zone (Patten 2003) 

[4] a—1-hr post treatment within treatment zone under canopy; b—1-hr post treatment, average of 3 samples (75% interception rate); c—geometric mean of all 

samples collected 3.5 to 4 hours post treatment (Patten 2003) 

[5] a—Empirical measurements from Hoerger and Kenag (1972) and Fletcher et al. (1984); b—from empirical measurements of Fletcher et al. (1984); c—from

modeled results of Hoerger and Kenaga (1972) 

4.1.4 Sediment Concentrations 

Limited testing of marine sediment concentrations following imazapyr treatment has been

conducted by Patten (2003), as described in section 3.0. The worst case value in sediment is

represented by the maximum sediment concentration detected in treated bare mud (5.7 mg/kg), the

upper limit would be represented by the average of the maximum values from Patten’s trials, and 

the typical environmental concentration presumes canopy interception, or 1.42 mg/kg (Table 4-1). 

All of these values are conservative in that the measurements were taken after the first tidal wash,

and hence represent “acute” sediment conditions as opposed to more chronic sediment conditions. 

The half-life in estuarine sediments will be substantially less than the 12.2-day half-life determined

in freshwater pond sediments by Mangels and Ritter (2000) because of the tidal exchange of

waters. However, due to the non-static nature of the estuarine environment, true sediment halflives cannot be determined from empirical measurements and “dissipation” rates more accurately 

describe what is actually occurring in the estuarine environment—capturing the multiple 

mechanisms that reduce sediment concentrations over time. Patten (2002) projects that the

sediment concentration of imazapyr after bare-mud treatment followed an exponential decline and 

could be predicted from the equation f = 0.0013 e((137/(x+23))

. In those studies, approximately one

fourth of the maximum detected concentration of imazapyr in sediment one hour after treatment

(5.7 mg/kg) was detectable after roughly 4 days post treatment (Figure 3-9), and the decay 

equation predicted the complete dissipation of the herbicide from sediment in 400 hours. 

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4.2 Ecological Receptor Exposures 

In this section, specific doses are estimated for imazpyr to terrestrial and aquatic receptors, by a

variety of exposure pathways of relevance to the habitats where Spartina treatments could occur.

Life history data for terrestrial wildlife were acquired from the Wildlife Exposure Handbook 

(USEPA 1993) and from anonymous (2003) for the bobwhite quail, marsh wren, deer mouse,

cottontail rabbit, mallard duck, Norway rat and lesser scaup. These species, or closely related 

species, utilize habits where Spartina is distributed, and/or they are test species for which 

toxicological data have been developed. The marsh wren is primarily a wetland and salt marsh

species that consumes a diet almost exclusively of animals and insects, and is particularly common 

in coastal areas. The deer mouse consumes a diet of seeds and nuts primarily and is found in a wide

variety of habitats throughout the state. The mallard is a “puddle” duck that consumes primarily 

vegetation, but also some invertebrates. The scaup is a “diving” duck and is more omnivorous in 

its dietary habits than the mallard. Both duck species will consume more high protein/fat animal

foods prior to periods of migration and breeding (Cohen et al. 2000). The cottontail rabbit is an

obligate herbivore and a test species used commonly to address dermal sensitivity. The Norway 

rat is a representative omnivore common to coastal areas, and the rat is also a commonly used test 

species. Average weights, surface areas, and daily consumption rates were used to represent 

exposure to wildlife species. These numbers can exhibit a great deal variation among populations,

but population-specific data from each of the areas where Spartina is distributed was not available.

Acute Dietary Exposure Modeling Method 

The acute dietary exposure was determined from the modeled empirical plant residue studies

discussed in section 4.1, taking the average plant residue detected from grassland applications of

herbicide at 1.5 lb ae/acre. Dose was modeled by the method of SERA (1999) used to address the 

ecological risks of imazapyr use in forestry applications for the USDA. Briefly, dose (D) = A x 

C(Prop)/W, where A = food consumption per day (kg), C = concentration in food (mg/kg), Prop = 

proportion of diet as treated vegetation (percent), and W = average body weight (in kg). Typical

and upper food concentration limits were obtained from residue studies outlined in Table 4-1, as 

previously discussed. Body weight and food consumption parameters were obtained from the

Wildlife Exposure Handbook or anonymous (2002) and are presented in Table 4-2. No dissipation

or degradation is assumed, and the acute exposure is presumed to equate to a “bolus” (single dose)

dietary exposure of the herbicide. The following parameters were assumed for this modeling: 

• Concentration in food: = 131 mg/kg (upper), 43 mg/kg (typical) 

• Proportion of Diet Contaminated (Prop): 

= 0.75 (obligate herbivore in treated Spartina meadow, based on 75% interception 

rate of Patten 2003) = upper limit exposure 

= 0.4 (based on 40% interception rate from Linders et al.2000) = typical proportion 

of diet assumed for omnivore and carnivore exposure. 

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Table 4-2: Exposure Parameters for Addressing Risks from WSDA’s Imazapyr Applications for Spartina Cont

Species 

Adult Body 

Weight 

(grams) 

Daily Food 

Intake 

(g) 

Daily Water 

Intake 

(ml) 

Inhalation 

Rate 

(m3/day) 

Surface 

Area 

(cm2) Diet Preference

Relevant Life History 

Characteristics Relevant to 

Exposure 

BobwhiteQuail 

B: 190 15 20 F: 0.10

M: 0.11 

F: 298 M: 320 

Plants and insects. Max insects20% in summer 

Breeding in April-July; hatchingMay to August;Non-migratory; annual mortality rate of approx. 80%

MarshWren 

B: 11.25 8 3 No data F: 45

M: 48 

Insects, spiders, mollusks, and crustaceans 

Breed in April; hatch in May; Migration in fall and spring; likely to be found within coastal marsh habitat where Spartina is abundant

Deer Mouse 

B:21 9

(lactating) 

7 F: .025

M: 023 

F: 86M: 91 

Mixture of nuts, seeds, and insects 

Breed several times during the year 

Cottontail Rabbit 

B: 1,286 180 125 0.63 1,254 Grasses, shrubs, woody plants Breed several times during the 

year 

Norway  Rat 

B: 300 15 33 No data 500 Omnivorous Breed several times during the 

year 

Mallard  Duck 

F: 1,043 M: 1,225 

250 F: 0.042

M: 0.055 

F; 0.42M: 0.48 

F: 1,030 M: 1,148 

A surface feeding “puddle” duck, feeds on an omnivorous diet. Dietary patterns vary withseason. In winter, mallards feedmostly on seeds mast, and to alesser extent invertebrates. Inthe migratory and breedingseasons, high protein and fatdiets are consumed, with moreinvertebrate biomass .

Red Fox F: 4,130 

M: 5,250 

F: 285 M: 362 

F: 355 M: 441 

F: 1.7 M: 2.0 

F: 2760M: 3220

Omnivorous: mostly small mammals, birds, insects, and fruit. Plant material is commonin summer and fall diet. 

Breeding in December - February 

FINAL October 2003 
Species 

Adult Body 

Weight 

(grams) 

Daily Food 

Intake 

(g) 

Daily Water 

Intake 

(ml) 

Inhalation 

Rate 

(m3/day) 

Surface 

Area 

(cm2) Diet Preference

Relevant Life History 

Characteristics Relevant to 

Exposure 

Scaup F: 770 

M: 860 

50 F: 0.064

M: 0.062 

F: 0.34M: 0.36 

F: 842 M: 906 

Juveniles ate entirely animal matter in NW territories study; 61% animal matter in Louisiana study, 

Pacific Flyway spring migration from March—April; fall migration from September-mid-October.

FINAL October 2003 
Chronic Dietary Exposure Method—On Site 

Chronic dietary exposure was evaluated using the same method adopted by the USFS in their 

evaluation of imazpyr use for forestry applications (SERA 1999), with several modifications

relevant to the proposed application rates to control Spartina. In the chronic dietary assessment, it

is assumed that consumption of treated vegetation will occur for a 90-day period beginning 

immediately after chemical application. Food consumption parameters are based on the Wildlife

Exposure Handbook reference as cited in Table 4-2. The residue of imazapyr in Spartina grain at

time zero (Co) is based on the acute dietary exposure scenario described above. The residue over

time was based on the foliar decay coefficient (k) = ln(2)/t50, where t50 = the foliar half-life. The 

concentration on the vegetation after time (Ct) is calculated as Ct = Coe

-kt

, and is reflected in

Figure 4-3 below. 

0

20

40

60

80

100

120

140

160

180

200

18

1522293643505764717885

Days Post Treatment

Estimated Pla

nt Re

sidue (mg/Kg)

Figure 4-3: Estimated plant residue concentrations over time, with an initial 

application rate of 1.5 lbs/acre.

The integral of the concentration after time t (Ct) is divided by the duration of dietary exposure to

calculate the time-weighted average concentration on vegetation. Daily dose is calculated from the 

proportion of the diet contaminated, divided by body weight (which varies by animal). The specific

parameters used are as follows: 

• Duration of exposure (T) = 90 days 

• Body weight (W)—average varies by animal, from Table 4-2, units in kg 

• Food consumption per day—varies by animal, from Table 4-2, units in kg 

• Foliar halftime (t50) = 37 days (upper limit), 26 days (typical exposure) 

• Foliar residue (rr): = 131 mg/kg (upper limit), 43 mg/kg (typical exposure) 

• Drift = 1.0 

• Decay coefficient (k) = ln(2)/t50 = 0.0462/day (upper limit); 0.0267/day (typical exposure) 

• Initial Concentration on Vegetation (Co) = Application Rate x (rr) x Drift

= 196.5 (upper); = 64.5 (typical) 

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October 2003 Page 65
• Concentration on Vegetation at time T (Ct) = Co e-kT

• Time-weighted average concentration on Vegetation (Ctwa) = Co(1-e-kt)/(kT) 

= 46.5 mg/kg (upper); 24.4 mg/kg (typical) 

• Proportion of Diet Contaminated (Prop): 

= 0.75 (obligate herbivore in treated Spartina meadow) 

= 0.4 (omnivore on meadow fringe, or transient wildlife) 

• Chronic Dietary Dose Absorbed (CD) = (Ctwa)(food intake/day)(Prop)/W 

Drinking Water Consumption Modeling 

Exposure to imazapyr through drinking water is unlikely due to the brackish water conditions 

where Spartina is found. It is possible, however, that small animals and insects will collect 

drinking water from the Spartina canopy after a rain. Thus, the drinking water pathway is

considered complete for the terrestrial wildlife considered in this assessment. 

Spill Scenario 

For this exposure scenario, the upper limit of exposure was taken to be drinking water obtained 

from the applied (undiluted) solution that could accumulate on the plants and be consumed (e.g., 

via licking). The concentration of imazapyr in the undiluted solution will vary by the spray

volume, as depicted in Figure 4-4. As demonstrated in this figure, the ultra-low spray volumes

result in the highest “neat” concentration of imazapyr, whereas high spray volumes dilute the

imazapyr in solution. Most application techniques envisioned would apply the herbicide in the 10 

gal/acre to 40 gal/acre range. Exposure via this method would be consistent with an accidental spill

scenario. Dose (D), in units of mg/kg-bw, is calculated as the product of (C x A)/W, where C = 

0

10000

20000

30000

40000

50000

60000

70000

80000

0 20 40 60 80 10

Application Volume (gal/acre)

Imazpyr Concentration in 

Applied Solution

0

Figure 4-4: Estimated concentration of undiluted imazapyr in various spray volume 

applications. 

Note: Higher spray volumes could be used and would result in lower exposures, as predicted. 

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October 2003 Page 66
concentration in ambient water (mg/L), A = water consumed per day (in liters), and W = body 

weight (in kg). This dose calculation was adopted from SERA (2001), with modifications to 

reflect the estimated exposure concentration. Body weight data for each of the animals are

represented in Table 4-2. For the exposure calculations, the upper limit exposure concentration 

(C) for the drinking water (spill) scenario presumed a spill (and ingestion) of concentrated spray 

solution that was to be applied at a rate of presumed to 10 gal/acre (17,966 mg-imazapyr/L), and 

the typical exposure was presumed to occur from a spill at the standard application rate of 20 

gal/acre (8,983 mg-imazapyr/L). 

Runoff scenario 

Ingestion of contaminated water from an acute exposure scenario for runoff represents a more

realistic case for what might actually occur in the treated environment. Again, however, due to the

brackish conditions where Spartina is found, it is unlikely that significant drinking would occur by 

any small mammal. Salt glands in ducks enable their ingestion of brackish water and for this

reason, an estimation of uptake from a runoff concentration is legitimate. The calculation of dose

is consistent with the equation used to predict uptake from a spill scenario (D = CxA/W), and the 

estimated environmental concentrations used for modeling are taken from the empirical results 

summarized in Table 4-2 (Patten 2003). Thus, the upper limit exposure was based on an EEC of 

3.4 mg-imazapyr/L, and the typical exposure was based on an EEC of 0.1 mg/L. 

Dermal (Contact) Exposure Modeling Method 

The ‘worst case’ (upper limit) scenario for contact exposure was based on the presumption of an 

animal being exposed directly to the spray, with 100% absorption ocurring over the first 24-hour

period. This is a highly conservative exposure model, in that it assumes 100% absorption of the

dose, which would overestimate fur bearing animal and bird exposures due to fur and feather

interference, respectively. Also, the dermis is largely impermeable to water soluble substances, 

thus, although adjuvants delivered with the herbicide will facilitate uptake, absorbtion of herbicide 

that reaches the skin will not be complete. The assumptions for the dermal absorbtion model were 

as follows: 

• Period of exposure (T) = 24 hrs 

• Body weight (W) = average weight in kg, as indicated from Wildlife Exposure Handbook, 

or other literature if no data were available within the exposure handbook. 

• Exposed surface area (SA): cm2 = 1110(W)0.65

• Application Rate (AR) = (1.5 lbs/acre)(0.01121—a conversion factor) = 0.0168 mg/cm

2

• Amount deposited on the animal (Amnt) = 0.5 x SA x AR (note: typical dermal exposure

assumes 50% of animal is covered by direct spray, upper limit exposure presumes 75% of 

animal is covered and the product of SA x AR is multiplied by 0.75. 

• Estimated Absorbed Dose = Dabs = Amnt/W

Other Exposure Pathways 

Additional routes of exposure are acknowledged both in the conceptual model (Figure 2-1) and 

also in Table 4-2. However, we consider the above exposure scenarios to represent the pathways

where exposure could be maximal. Pathways such as chronic exposure to contaminated water are

incomplete for the proposed use of Spartina in an estuarine setting. The acute run-off scenario to

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contaminated water—where an animal might drink water that has been in contact with treated 

vegetation is also unlikely to be completed in the estuarine setting because the brackish waters

where Spartina is distributed are not potable to most wildlife (although waterfowl with salt glands 

are capable of drinking a limited amount of salt water). We did not model this type of exposure, 

as the more severe exposure scenario, the acute spill based on the drinking of neat solution,

addressed a more significant exposure. 

Acute inhalation exposure is possible, but considered insignificant because the disturbance created 

during treatment will cause most animals to avoid the area being immediately treated. Aerial drift 

will vary by nozzle type and application volume, and has been estimated at 10% in grasslands in 

the Netherlands (Linder et al. 2000). Such drift can be both a component of inhalation exposure

off-site, and/or oral or dermal exposure off-site. The probability of this occurrence representing

substantive exposure is considered insignificant, and the exposure modeling conducted for on-site 

exposures will yield substantially higher doses than potential exposures off-site. Since these

results indicated insignificant risk (as described below), off-site exposure from drift was not

modeled. 

4.2.1 Mamalian Exposure 

Mammalian wildlife can be exposed to imazapyr through dermal, oral (ingestion) or inhalation

routes, although the dietary route is considered most likely in practical settings (USDA 1996a). We

modeled exposure to the rabbit, rat, mouse and fox as representative mammalian herbivores and

omnivores for those areas where imazapyr could be applied to control Spartina. As demonstrated in

Table 4-3, the acute dermal and dietary exposures yielded the highest modeled doses to all of the

mammalian wildlife examined for this study for those pathways that are most realistic in-situ (i.e., 

not involving a spill scenario) Based on life history characteristics of food intake, body weight, and

surface area, the deer mouse would have the highest estimated exposure of the mammalian 

ecological receptors considered, for all of the exposure pathways considered. This finding is

relatively intuitive in that the species has the smallest home range, and highest food and water 

intake and surface area relative to body weight. Estimates of dose within each exposure pathway 

varied by slightly more than one order of magnitude amongst the mammalian species examined 

(Table 4-3). The red fox had the lowest modeled exposure from contact exposure, whereas the

Norway rat had the lowest modeled exposure for the acute and chronic dietary scenarios. The 

smaller surface area/body weight ratio of the fox, and the omnivorous habits of both the fox and rat,

were largely responsible for the lower estimated exposures by these species, relative to the mouse

and rabbit. 

Estimates of imazapyr exposure ingested from an acute spill scenario were high for all mammalian 

species, yielding acute dose estimates in the high mg/kg-bw to low g/kg-bw range, for the upper and 

typical spill concentration estimates. These acute “drinking water/spill” scenario values exceeded

the estimated exposure for other pathways by two to three orders of magnitude. However, it is

likely that all of the mammalian species modeled would voluntarily avoid drinking water containing

imazapyr from a spilled, undiluted spray solution; thus, the practical use of these modeled exposures 

requires field validation. Drinking water consumption under the run-off scenarios yields more 

realistic estimates of exposure, and reduced the exposure doses to the modeled mammals by over

three orders of magnitude, with a maximum estimated exposure in the deer mouse of 1.133 mg/kg, 

and a minimum in the male fox of 0.008 mg/kg—the lowest dose estimated by all exposure routes

considered. 

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Table 4-3: Estimated Exposures to Terrestrial Wildlife Receptors from Imazapyr Applications (mg/kg-body wt

Receptor Acute Dietary Exposure

Chronic Dietary

Exposure 

Acute Drinking Water 

Exposure (Runoff) 

Acute Drinking Water 

(Spill) Exposure

Deer Mouse Upper: 42.11 

Typical: 13.82 

Upper: 14.95 Typical: 7.84

Upper: 1.133 Typical: 0.033

Upper: 5 989 Typical: 2 994 

UT

Scaup Upper: 3.05 M, 3.40 F 

Typical: 1.00 M, 1.12 F 

Upper: 1.08 M, 1.21 F Typical: 0.57 M, 0.63 F 

Upper: 0.210 M, 0.216 F Typical: 0.006 M, 0.006 F

Upper: 1107 M, 1143 F Typical: 554 M, 572 F 

UT

Mallard Duck Upper: 20.05 M, 6.58 F 

Typical: 6.58 M, 7.73 F 

Upper: 7.12 M, 8.36 F Typical: 3.73 M, 4.39 F 

Upper: 0.186 M, 0.143 F Typical: 0.004 M, 0.005 F 

Upper: 983 M, 758 F Typical: 491 M, 379 F 

UT

Norway Rat Upper: 2.62 

Typical: 0.86

Upper: 0.49 Typical: 0.93

Upper: 0.374 Typical:0.011 

Upper: 1 976 Typical: 988 

UT

Marsh Wren Upper: 37.26 

Typical: 12.23 

Upper: 13.23 Typical: 6.94

Upper: 0.903 Typical: 0.027

Upper: 4 7910 Typical: 2 395 

UT

Bobwhite Quail Upper: 7.757 

Typical:2.546 

Upper: 2.75 Typical: 1.44

Upper: 0.358 Typical: 0.011

Upper: 1 891 Typical: 946 

UT

Cottontail Rabbit Upper: 13.75 

Typical: 4.51

Upper: 4.88 Typical: 2.56

Upper: 0.330 Typical: 0.010

Upper: 1 746 Typical: 873 

UT

Red Fox Upper: 3.61 

Typical: 1.19

Upper: 1.28 Typical: 0.67

Upper: 0.286 M, 0.292 F Typical: 0.008 M, 0.009 F 

Upper: 1509 M, 1544 F Typical: 755 M, 772 F 

UT

*See Appendix B for full calculations of dose by exposure pathway for each receptor species modeled.FINAL October 2003 

The lowest dietary dose was modeled in the rat at 0.049 mg/kg, for a “typical” chronic dietary dose

(see Table 4.2). The highest dietary dose was calculated for the acute dietary exposure in the deer 

mouse consuming a diet with the “upper” limit EEC, to yield a dose of 42.11 mg/kg. The highest

non-spill dose was estimated in the deer mouse via direct contact to spray, yielding a dose estimate

of 54.07 mg/kg-bw. 

4.2.2 Avian Exposure 

Exposure for birds may occur via the same pathways as mammals: ingestion, contact and 

inhalation. The broad array of life history behaviors prevents an assessment of all bird species.

We modeled potential doses to the bobwhite quail, marsh wren, mallard duck and greater scaup, to 

provide for a range of dietary habits and life history behaviors. All of these species, with the 

exception of the bobwhite quail, could be found in the regions of Washington State where Spartina

is currently distributed (western Washington’s lowlands support California quail, a closely related 

species). As demonstrated in Table 4-3, the marsh wren had the highest projected dose among 

the species via the non-drinking exposure pathways considered (i.e., 67 mg/kg via direct contact 

application). The highest acute and chronic dietary doses were also projected for the marsh wren

of all the bird species, primarily due to its high food intake relative to its body weight. Similarly,

the mallard duck was estimated to consume roughly six-fold more imazapyr than the greater scaup 

because of the assumed dietary differences considered between these species (i.e., primary 

herbivore vs. omnivore). 

4.2.3 Insect Exposure 

Exposure for insects was modeled in the bee for the direct contact exposure scenario due to spray 

from an acute exposure only (Meade 1983). Dose was estimated as described in Section 4.1 for

direct contact exposure, using the bee as a surrogate for all terrestrial insects. The direct contact 

exposure for the bee yielded an estimated dose of 0.0223 mg/kg for a typical exposure (assuming 

50% of the insect was covered), and 0.0335 mg/kg for the upper limit estimate (assuming 75% of

the animal was covered). Dietary and drinking water consumption were not considered, as

previously discussed. 

4.2.4 Reptiles and Amphibian Exposure

Reptiles and amphibians (herptiles) can be exposed to herbicides through the same pathways as 

mammals, birds and terrestrial invertebrates—via dietary consumption, inhalation and direct

contact. Amphibians may be particularly susceptible to contact exposure from direct spray

applications whereas contact exposure to reptiles is unlikely to yield significant doses due to the 

relative impermeability of their skin. Exposure parameters have not been developed to accurately 

gauge reptile or amphibian exposures and no toxicity information was identified from the literature 

from which to quantify or estimate exposure to these groups of animals. 

Although a formal dose cannot be estimated for herptiles, the potential for exposure can be

considered by evaluating the potential herptile inhabitants of the estuarine areas where Spartina

treatments could occur. The USGS (2003b) reports a total of 19 herptiles potentially resident in the 

broader Willapa Bay area (http://www.npwrc.usgs.gov/resource/othrdata/chekbird/r1/willrept.htm). 

These species are also possible in other areas of Puget Sound and coastal Washington. Twelve of

these 19 species are amphibians and are therefore intolerant of the saline conditions inherent to 

estuaries where Spartina is distributed. Exposure to these 12 species is therefore considered an

incomplete pathway.

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October 2003 Page 71
Three of the 19 herptile species recognized to Willapa Bay are sea turtles, and include the

leatherback (Caretta caretta), loggerhead (Dermochleys coriacea) and green sea turtle (Chelonia

mydas). All of these species are listed as either threatened or endangered. Willapa Bay is not 

designated as critical habitat for any of these species (50 CFR Part 226), and we could not verify

that individuals of these species’ have been specifically sighted in the area. Documentation 

reviewed suggests very infrequent sightings of these sea turtles are possible in the outer bay and

marine areas due to their relatively cosmopolitan oceanic distributions. However, the green turtle

is considered primarily a tropical species whereas the loggerhead and leatherback turtles have a 

greater tolerance for colder waters and would be more likely to be observed off of coastal

Washington. Where Spartina is distributed, the probability of turtle occurrences is sufficiently low

to consider exposure an incomplete pathway for these threatened and endangered species. 

Only two snake species are recognized as typical inhabitants of the Willapa Bay area, the common 

gartersnake (Thamnophis sirtalis) and the northwestern gartersnake (Thamnophis ordinoides). 

These species may be also found elsewhere in coastal regions of western Washington

environments near where Spartina occurs. These species can be found occasionally in or near

water, although they would be considered very uncommon in salt marsh habitats. The 

northwestern garter snake is primarily a terrestrial species and the common garter is commonly 

found near freshwater which it may use to escape predation or other disturbance (Stebbins 1985). 

The potential for exposure to these animals is extremely low due to their life history behavior that

would largely preclude their use of salt marsh habitat where Spartina is distributed. No doses are 

estimated for these receptors. 

4.2.5 Aquatic Animal Exposure 

Exposure to fish and aquatic invertebrates could occur from drift entering water during application,

from runoff from plants following application from rain and tidal inundation, and/or from leaching 

from treated vegetation and sediment underlying treatment areas. The estimated water 

concentrations to which aquatic animals could be exposed will vary with depth following initial

application, as depicted in Figure 4-2. For the exposure calculations, the worst case exposure

concentration (C) for water was assumed to be 5.77 mg/L, the maximum concentration detected on 

the edge of the treatment zone on an incoming tide after application to bare mud (Patten 2003). 

The upper limit was assumed to be 3.40 mg/L, and the typical concentration was 0.1 mg/L, as

summarized in Table 4-1. Benthic infauna and epifauna atop mudflat sediments will be exposed to 

the highest concentrations. Sediment detritivores, benthic epifauna and benthic infauna were

assumed to be exposed to concentrations of 2.27, 1.42 and 1.4 mg-imazapyr/kg-sediment to 

represent the worst case, upper limit and typical concentrations found in sediment by Patten 

(2003). Benthic infauna were assumed to be exposed to the pore water levels detected in the Patten

study (2003), 3.29, 2.94 and 0.042 mg/L to represent the worst case, upper limit and typical

exposure scenarios. 

As previously discussed, the bioconcentration and bioaccumulation of imazapyr in aquatic 

organisms is extremely low due to the compounds high water solubility and low lipid solubility. 

Empirical results detailed in Table 4-1 of this document highlight how measurable concentrations

in fish and aquatic invertebrate tissues were detected only in samples collected 3 hours after 

treatment, but subsequent measurements were not detectable. Therefore the potential of exposure

through ingestion of exposed aquatic invertebrates or other food sources is substantially reduced. 

No bioaccumulated doses are therefore estimated. 

FINAL 

October 2003 Page 72
4.3 Adjuvant and Inert Ingredient Exposure Assessment 

The mass of surfactant applied per unit surface area can change with the application volume used 

to dilute the herbicide. As a result, it is essential to consider the potential exposure to aquatic

animals at the range of potential application volumes. A variety of surfactants could be used, on 

the basis of past practices, but all are assumed to be mixed with herbicide at a rate of 1% (w/v). 

We evaluated the potential concentration of surfactant over a range of potential application 

volumes used, ranging from the ultra-low range (2.5 gal/acre) as applied in some treatment trials

by Patten (2002) to 80 gal/acre, which is consistent with the rates used to apply glyphosate under 

the current chemical control program (WSDA 1993). As demonstrated in Table 4-2, initial

concentrations of surfactant could be projected to be quite high under the most conservative of

assumptions and highest application rates. At a water depth of 0.0125 m (approx. 1⁄2 in), as would 

be consistent with an incoming tidal lens, the spray volume of 80 gal/acre could be projected to

yield nearly 60 mg/L surfactant, whereas low and ultra-low rates of application yield 

correspondingly lower concentrations. 

Table 4-4: Estimated surfactant concentrations in estuary waters where Spartina could be 

treated with imazapyr, assuming constant surfactant rate of 1% in spray volume, complete

solubility on incoming tide, and the “worst case” scenario of no adsorption on sediment or

Spartina canopy. Surfactant concentrations are in mg/L 

Depth (m) 

2.5 

gal/acre 5 gal/acre 10 gal/acre 20 gal/acre 40 gal/acre 80 gal/acre 

0.0125 1.872 3.744 7.48 14.96 29.92 59.84

0.025 0.936 1.872 3.74 7.48 14.96 29.92

0.05 0.468 0.936 1.87 3.74 7.48 14.96

0.1 0.234 0.468 0.935 1.87 3.74 7.48

0.2 0.117 0.234 0.468 0.935 1.87 3.74

0.4 0.058 0.117 0.234 0.467 0.935 1.87

0.8 0.029 0.058 0.117 0.234 0.468 0.935

FINAL 

October 2003 Page 73
5.0 RISK CHARACTERIZATION TO ECOLOGICAL RECEPTORS 

FROM THE USE OF IMAZAPYR TO CONTROL SPARTINA

To characterize risks from the potential practice of applying imazapyr in an estuary setting, the

potential imazapyr exposure and effects to ecological receptors are integrated in a total ‘weight of

evidence analysis’. The analysis examines the estimated dose and chemical properties (toxicity,

metabolism, environmental fate and transport) that influence the impact that imazapyr could have 

on biota. Characterization of risk was made by calculating the hazard quotient from the estimated

environmental concentrations or exposure doses calculated in section 4.0 of this report. The hazard 

quotient is derived by dividing the estimated exposure concentration or dose by the toxicity

reference dose (NOEL) reported for the species (or closest surrogate species) for which toxicity

testing has been conducted (Table 5-1). When possible, we used the NOEL derived for the same

exposure pathway for which a dose was estimated to calculate ‘pathway-specific’ hazard quotients. 

When pathway-specific NOEL’s were not available, we used the most conservative NOEL from 

the literature for all of the exposure pathways considered, defaulting to the one-compartment

model assumption for uptake and elimination kinetics. 

Based on past toxicological research and the physical chemistry of imazapyr, it is assumed that

uptake of imazapyr into an animal follows a first compartment model with first-order elimination 

kinetics. That is, it is assumed that once exposed, the imazapyr does not preferentially concentrate 

in one tissue over another, and that it is eliminated from all tissues at effectively the same rate.

Under a one-compartment model, uptake of imazapyr by one pathway will have an equivalent 

effect as the same dose acquired by another pathway. Thus, the most conservative toxicological 

criteria would be applicable to all means of exposure. However, in practice, even with a onecompartment conceptual model, this scenario is rarely observed and differences in the doses

required to elicit a toxicological response are often observed among different exposure pathways. 

For example, skin barriers may prevent full absorption of a dermally applied dose, or portions of an 

ingested quantity of herbicide may pass through an animal’s system without systemic absorption in 

the small intestine. 

5.0.1 Mammal Risk 

As discussed in chapter 3 the acute oral toxicity of imazapyr to mammals is rated as practically

non-toxic, based on the EPA criteria outlined in Table 2-1 (i.e., an acute oral LD50 of > 2,000, 

mg/kg-body weight). None of the exposure doses estimated in chapter 4 (see Table 4-3) exceeded a

hazard quotient of 1 for any of the species or exposure pathways modeled relative to the NOEL

(Table 5-1), with the exception of the deer mouse spill scenario exposure (HQ = 1.198). The spill

scenario modeled (i.e., where an animal would effectively drink undiluted spilled spray solution) is

highly conservative and unlikely to be realized in situ because best management practices would 

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Table 5-1: Hazard quotient calculations from estimated exposure doses of imazapyr to terrestrial wildlife.* 

Ecological 

Receptor 

Acute 

Dietary 

Upper 

Exposure1

Acute 

Dietary 

Typical 

Exposure1

Chronic 

Dietary Upper

Exposure2 

Chronic Dietary

Typical 

Exposure2 

Dermal/ 

Contact Upper 

Exposure3 

Dermal/ 

Contact 

Typical 

Exposure3 

Drinking 

Water (Spill) 

Upper 

Exposure4

Drinking Water 

(Spill) Typical 

Exposure4

Deer Mouse 0.0084214 0.000524 0.014946429 0.007842857 0.02703254 0.0180217 1.19773333 0.59886667 Norway Rat 0.000524 0.000172 0.00093 0.000488 0.01065789 0.0071053 0.395252 0.197626 Cottontail Rabbit 

0.0027504 0.0009028 0.012203538 0.006403577 0.13516268 0.0901085 0.34926128 0.17463064 

Red Fox M 0.0007226 0.0002372 0.005130057 0.002691901 0.05328943 0.0355263 0.3018288 0.1509144 Red Fox F 0.0007232 0.0002374 0.00513414 0.002691901 0.03201837 0.0213456 0.3088586 0.1544293 Marsh Wren 0.0552852 0.018147 0.066133333 0.034702222 0.01956941 0.0130463 7.07675744 3.53837872 Bobwhite Quail 0.0115083 0.0037775 0.013766447 0.007223684 0.02128389 0.0141893 2.80587225 1.40293612 Scaup M 0.0152326 0.0008703 0.005406977 0.002837209 0.16816289 0.1121086 0.96362661 0.48181331 Scaup F 0.017013 0.000972 0.006038961 0.003168831 0.06252731 0.0416849 0.99503125 0.49751563 Mallard Duck M 0.1002551 0.0057281 0.035586735 0.018673469 0.03686032 0.0245735 0.8552047 0.42760235 Mallard Duck F 0.1177493 0.0067277 0.0417965 0.021931927 0.03831438 0.0255429 0.65962899 0.3298145 

1: acute dietary reference NOEL = 5,000 mg/kg-bw for mouse, rat, rabbit, and fox; 1149 mg/kg for mallard and scaup; and 674 

2: chronic reference NOEL dose for mouse and rate = 1,000 mg/kg; for rabbits = 400 mg/kg; for dogs = 250 mg/kg; for avian s3: dermal/contact NOEL dose for mouse & rat= 2,000 mg/kg; 400 mg/kg for rabbit. No data on fox or avian species, so assum4: same reference doses as acute oral 

FINAL October 2003 

be employed immediately to clean up spilled herbicide, and the disturbance of the cleanup action

would discourage wildlife from the area. All other exposure scenarios and estimated doses for the

terrestrial mammals modeled yielded hazard quotients two to four orders of magnitude lower than

the NOEL values for the species or guild from which toxicological data were derived. This results 

indicates insignificant risk to these receptors can be expected from imazapyr treatments. 

Characterizing risk based on absolute lethal thresholds such as the LD50 is not possible for

mammals because the dose ranges administered over the variety of tests performed have never

yielded lethality in mammals. Thus, the estimates of the LD50, as reported in Table 3-9, are 

prefaced with the ‘>’ sign, and no empirical results are available to substantiate these values. 

Substantial conservatism was factored into the exposure assessment such that the modeled doses

can be assumed to overestimate the conditions in-situ, particularly for chronic exposures because

applications will occur only every other year and tidal flushing over the Spartina results in the loss

of the herbicide over time. Conservatism was also factored into the hazard quotient calculations, in

that most NOELs reported simply referenced the highest dose tested (HDT), and therefore were not

based on actual empirical findings from a dose-response curve. It is therefore the conclusion of 

this assessment that the use of imazapyr at the indicated application rate of 1.5 lbs/acre (0.68 

kg/acre) will not pose a risk to mammalian ecological receptors, even under the worst case (upper)

exposure scenarios. No threatened or endangered mammalian species occupy habitats where

Spartina is distributed or where imazapyr could be applied. Since the chemical does not 

bioaccumulate, and best management practices will prevent significant drift off-site, it can be 

reasonably assumed that such species occurring off-site would not be affected by the use of 

imazapyr in the estuary setting. 

5.0.2 Avian Risk

As discussed in chapter 3 the acute oral toxicity of imazapyr to birds is also rated as practically 

non-toxic, based on the EPA criteria outlined in Table 2-1. None of the exposure doses estimated 

in chapter 4 (see Table 4-3) yielded a hazard quotient of 1 or greater for any of the species or

exposure pathways modeled relative to the NOEL except, again for the spill scenario where both

the marsh wren and bobwhite quail had upper limit exposure HQs of approximately 7 and 3, 

respectively, and lower limit HQs of 3.5 and 1.4 (Table 5-1). The same argument holds true for

avian wildlife as it would for terrestrial wildlife regarding the spill scenario—namely that best

management practices that obligate WSDA to immediate clean-up actions create disturbance that

would be expected to greatly eliminate exposure to birds of this guild when herbicide

concentrations are at their highest. The more realisitic exposure scenarios-- run-off drinking water,

dietary and direct contact yielded hazard quotients generally two to three orders of magnitude 

below 1, indicating insignificant risk to these receptors can be expected. 

Similar to mammals, characterizing risk based on lethal thresholds is not possible for birds. Like 

mammals, in the toxicity tests conducted where survival was a measurement endpoint, the dose

ranges administered did not yield toxicity (lethality), so the estimates of the LD50, as reported in

Table 3-10, are prefaced with the ‘>’ sign, and no empirical results are available to substantiate

these values. Substantial conservatism was therefore factored into the exposure assessment such

that the modeled doses can be assumed to overestimate the conditions in-situ, particularly for 

chronic exposures. By considering avian species with a range of life history behaviors and dietary 

habits, we have screened for a range of avian receptors that could be exposed to imazapyr in the 

environments where it might be applied. It is the conclusion of this assessment that the use of 

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imazapyr at the indicated application rate of 1.5 lbs/acre will not pose a risk to avian receptors, 

even under the worst case exposure scenarios. 

5.0.3 Insects 

Imazapyr is practically non-toxic to honey bees (BPA 2000). Direct contact exposure resulted in 

an estimated dose of 0.0223 mg/kg and 0.0335 mg/kg for typical, and upper limit exposure, 

respectively. The estimated NOEL for insects is 1000 mg/kg (the HDT), and the LD50 is 

considered > 1,000 mg/kg. On this basis, the hazard quotients would be 0.000223 and 0.000335, 

respectively, and the risks can be characterized as insignificant to terrestrial insects, even from the

“worst case” contact exposure scenario. 

5.0.4 Reptiles and Amphibians 

No imazapyr toxicity values have been reported for reptiles and amphibians for imazapyr, and 

exposure parameters have not been fully developed, as discussed in Section 4. Although a formal

dose (and hence risk calculation) cannot be extrapolated, the life history behaviors of the herptiles 

native to western Washington suggests that risk for reptiles and amphibians is insignificant because 

they would not be found in the brackish water and estuarine habitat where Spartina could be

treated with imazapyr, and thus exposure is precluded, as described in section 4.2. 

5.0.5 Fish 

Risk to aquatic ecological receptors from exposure to a potentially hazardous substance is 

determined by two equally important factors: duration of exposure and the concentration or dose

of the chemical (which is a function of the potency or toxicity of the chemical).The acute toxicity 

(LC50) concentration for rainbow trout, bluegill sunfish, and channel catfish was formerly reported 

at >100 mg/L (the HDT), and as such the herbicide has been considered practically non toxic

based on EPA hazard criteria (Table 3-12) . As discussed in this assessment, more recent toxicity 

testing has established an empirical LC50 of 22,305 mg-imazapyr ae/L using the Arsenaltm

formulation. This toxicity value actually exceeds the imazapyr concentration in a neat (undiluted)

spray solution applied at the normal 10 gal/acre application rate (i.e., 17,966 mg-imazapyr/L), and 

is within one order of magnitude of the neat concentrations if applied at the ultra-low rates of 5 or 

2.5 gal/acre (35,931 and 71,862 mg/L, respectively). Based on the worst case, upper limit and

typical exposure concentrations outlined in chapter 4.0 (Table 4-1), as developed from empirical 

studies, the hazard quotients for water exposures to rainbow trout would be 0.00258, 00152, and 

0.000045, respectively (Table 5-2). These values represent insignificant risk to fish. 

It should also be understood that these exposures are relevant only for an acute exposure scenario. 

No significant chronic exposures should occur due to the tidal exchange of waters in the areas 

where Spartina is distributed. Based on the dissipation experiments of Patten (2002), all readily 

solubilized imazapyr applied during a typical application would be dissipated (diluted beyond 

detection) in approximately 40 hours or less—roughly four to five tidal exchanges in the coastal

waters of Washington. This duration of exposure is less than half the time of a typical 96-hr 

toxicity test. 

Risks to Threatened and Endangered Fish 

Based on the hazard quotient calculations described above, and the threatened and endangered 

species sensitivity fish toxicity study recently conducted (Sappington et al. 2000), it can be 

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reasonably assumed, with high certainty, that exposure to the active ingredients in imazapy pose no 

significant risk to the Columbia River ESU chum salmon, chinook salmon, or candidate coho 

salmon stocks, nor to any of the Puget Sound salmonid stocks that are listed as threatened or

endangered under the ESA. Only under the accidental spill scenario of undiluted neat solution that

was to be applied at a rate of 2.5 or 5 gal/acre is there potential to exceed the acutely toxicity 

criteria for salmonids (as active ingredient). However, if a spill were to occur in an aquatic 

environment where salmon were inhabitants (i.e., as opposed to a bare mud-flat at low tide), water

must be present. Thus, some dilution would occur immediately and the dilution would likely lower

the concentration below the toxicity threshold. 

5.0.6 Aquatic Invertebrates 

The reported acute toxicity LC50 concentration for the water flea Daphnia magna is >100 mg/L 

and the reported acute toxicity LC50 concentration for the eastern oyster growth inhibitation is > 

132 mg/L (see section 3.1.9). On the basis of these toxicity measurements, imazapyr would be

considered practically non-toxic to both freshwater and marine invertebrates according to EPA

hazard screening criteria (Table 3-11). However, similar to mammal studies, and most fish studies

(beyond those recently conducted by Smith et al. 2003) it should be recognized that no formal

measurements have been documented that establish imazapyr concentrations that were empirically 

toxic or yielded sub-lethal effects such as growth inhibition. Thus, the measures of > 100 and > 

132 mg/L provide only screening values for this current effort. To differentiate risks from motile

epibenthic or pelagic invertebrates from benthic infauna we used the pore water concentrations

depicted in Table 4-1. No sediment quality data have been derived to enable risk characterization 

from potential sediment contamination, although epibenthic invertebrates such as the oyster and 

crayfish have been tested. Using these toxicity measures and the estimated worst case, upper limit

and typical exposure concentrations from Table 4-1, the hazard quotients have been summarized in 

Table 5-2. 

As depicted in Table 5-2, conservative risk modeling assuming acute exposure conditions most 

likely to be experienced by invertebrates yielded hazard quotients two to three orders of magnitude

below 1 for the active ingredient of imazapyr. These results suggest that invertebrate exposure to 

imazapyr in water represents an insignificant risk. 

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Table 5-2. Hazard Quotient Calculations from Estimated Aquatic Biota Exposures* 

Species Worst Case Exposure Upper Limit Exposure Typical Exposure 

Rainbow Trout 

(LC50 = 22,305 mg/L) 

0.00258 0.00152 0.000045

Rainbow Trout, Arsenal (w/1%

surfactant) 

a) Hasten 113 mg/L LC50 

b) W/Agridex: 459 mg/L

LC50 

a) 0.511 

b) 0.126 

a) 0.301 

b) 0.074 

a) 0.0088 

b) 0.0021 

Marine Invertebrates (surface

water exposure, NOEL = 100

mg/L) 

0.0577 0.034 0.001

Marine Benthic Infauna

Invertebrates (pore water

exposure; NOEL = 100 mg/L) 

0.0329 0.0294 0.00042

Non-target algae (e.g., Ulva 

sp., etc.—EC50 = 71 mg/L in

Selenastrum) 

0.076 0.0447 0.0013

Non-target vascular plants 

(e.g., eelgrass, Salicornia; 

EC50 =0.0214 mg/L in

duckweed for growth) 

270 159 4.7

*Hazard quotient defined as: estimated exposure concentration/sub-lethal EC50, or estimated exposure concentration/LC50

5.0.7 Non-target Aquatic Vegetation Risk 

It is not surprising that risks to non-target aquatic vegetation appear to pose the most significant

risk element from the potential use of imazapyr, as the herbicide has been engineered as a broadspectrum agent to control unwanted plant growth. Risks to algae, based on hazard quotient

calculations, are insignificant—within the same order of magnitude as risks characterized for

aquatic invertebrates (Table 5-3). Risks to vascular plants such as eelgrass (e.g., Z. marina and Z. 

japonica) may be significant, based on EC50 concentrations developed in duckweed, a floating

vascular plant and expected water concentrations of the herbicide. Hazard quotients exceeded 1.0 

under each of the exposure scenarios considered for vascular plants, ranging from 4.7 fold above 

the EC50 for duckweed growth inhibitation for typical exposures to 270 for “worst case” exposures.

The impact of imazapyr use on non-target vegetation should be largely controllable by the use of 

best management practices that limit the potential for non-target vegetation exposure. Patten 

(2003) showed that even after direct exposure to the herbicide, regrowth of eelgrass to its preapplication state was evident less than one year after treatment. This effect was observed in 

eelgrass specifically treated with the herbicide, not treated through exposure to water as is

considered the most realistic means of exposure in this context. The monoculture growth typical of

Spartina reduces the potential for non-target plant exposure during herbicide application. 

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5.1 Relative Hazard and Risk Characterization of Imazapyr, Glyphosate, 

Surfactants, Inert Ingredients, And The Use of No Chemical Control Or Other 

Agents For Spartina Eradication 

Glyphosate 

The full suite of ecological hazards and risks associated with glyphosate use to control Spartina

were thoroughly addressed in the original EIS (WSDA 1993). The conclusion of that document 

was that the use of the compound to control Spartina had limited risks associated with it to 

ecological receptors and therefore only those elements relevant to a comparison with imazapyr will

be evaluated here. On the basis of the multitude of factors evaluated in this assessment it can be 

summarized that imazapyr presents a substantially improved relative risk scenario to aquatic

animals over that of glyphosate. Reported LC50 values for glyphosate in rainbow trout range over

two orders of magnitude, from low values of 2 to 50 mg/L (Folmar et al. 1979, Hildenbrand et al. 

1982, Wan et al. 1989—as cited in Pan et al. 2002), to high values of > 1,000 mg/L (Geisy et al. 

2000). As presented in this report, the most recent (unpublished) data from EPA standard (1991)

96-hour static renewal tests of glyphosate (Rodeotm) with rainbow trout conducted concurrently 

with a comparison to imazapyr yielded an LC50 of 782 mg/L for Rodeo (95% confidence interval 

of 719-845 mg/L), relative to the LC50 of the Arsenaltm herbicide formulation of 77,716 mg/L 

(22,305 mg imazapyr/L) (Grue, C., personal communication 2003). Because these latter tests were 

conducted with fish from the same origin and age, in the same laboratory, the relative toxicity 

comparison is perhaps the most relevant of all the previous testing conducted. Based on these

results, the inherent aquatic toxicity of Rodeo may be approximately 28.5 to 99-fold more acutely 

toxic to fish than imazapyr—depending on whether the comparison is made to the active ingredient 

or the herbicide formulation, respectively. Using the more conservative toxicity estimates

established by the other researchers previously referenced, this difference could increase up to two 

orders of magnitude more than that developed from the data of Smith et al. (2002) and Grue (pers 

comm). The wide variation in glyphosate’s aquatic toxicity has been attributed to the dilution 

water, temperature, formulation, and the amount of suspended sediment in the water. Toxicity 

appears to increase with temperature, and decrease with elevated pH and suspended sediment

(WSDA 1993E, Schuette 1998). 

A similar comparison of the relative hazards to invertebrates between glyphosate and imazapyr

cannot be provided because no tests have been conducted with invertebrates at high enough 

concentrations of imazapyr to elicit mortality (i.e., NOEL and LC50 values represented were the

highest doses tested). However, the relationship between fish and aquatic invertebrate toxicity for 

a given chemical rarely differs by more than an order of magnitude (generally less than 1⁄2) so it is 

reasonable to expect a similar relationship to exist for the relative invertebrate hazards from 

glyphosate verses imazapyr. 

Glyphosate is an organophosphate compound, but it does not inhibit acetylcholinesterase activity 

like organophosphate insecticides such as Nuvantm or Diazinontm because the herbicide is missing 

an ester in its chemical structure (Pan et al. 2002). It elicits its herbicidal activity by inhibiting the

enzyme 5-enolpyruvylshikimate-3-phosphate synthase, which is needed by plants to synthesize

chorismate—needed in turn for protein synthesis in plants. Although highly soluble like imazapyr 

(solubility = 11,600 mg/L at 25o

C) with low potential for bioaccumulation, glyphosate—unlike

imazapyr—is not degraded significantly in water by photodegradation (Schuette 1998). It appears 

to adsorb readily to sediment, with half-lives on the order of 1.5 to 11.2 days, and sediment appears 

to be the principal environmental sink for this widely used herbicide. By comparison with 

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imazapyr, glyphosate has a low organic carbon partition coefficient indicating it can be expected to 

be substantially less mobile than imazapyr. In an estuary setting, the physical chemistry of 

glyphosate such as its lack of photodegradation and ready adsorption to sediment, suggests that

glyphosate will be more persistent than imazapyr. Recent results in Spartina treated meadows

have demonstrated that the rhizome concentrations of glyphosate increased 231 to 591% over a 

three year period, while sediment concentrations adjacent to the plots declined 88 to 96% (Kilbride

and Paveglio 2001). These recent results show that the residual biomass of Spartina could also

serve as a residual compartment of glyphosate for eventual release to the intertidal environment. 

Although significantly more toxic than imazapyr, the aquatic toxicity hazard rating of the herbicide

would still qualify as practically non-toxic to slightly toxic based on EPA toxicity criteria (Table 

3-12) and the species tested. (The terrestrial wildlife toxicity rates similar to imazapyr--practically 

non-toxic). However, at issue in the relative risk comparison is not simply the inherent aquatic 

toxicity of the compound, but also the estimated application rates relative to imazapyr. According

to recent results by Patten and Stenvall (2002), effective control with glyphosate was obtained only

with minimum application rates of 18 kg ae/ha, 10.7-fold greater than the 1.68 kg ae/ha needed for 

control with imazapyr. Thus, based on active ingredient concentrations required for herbicide

efficacy and inherent toxicity, the use of glyphosate reduces the margin of safety relative to 

potential environmental exposure concentrations by up to three orders of magnitude—irrespective

of the other associated risks from the surfactants required for application. 

Surfactants

The inherent risks of using either herbicide (i.e., imazapyr or glyphosate) rises significantly when 

mixed with surfactants, thus the choice of which surfactant is used is not trivial. In addition spray

volumes required differ by herbicide. Glyphosate applications require 80 to 100 gal/acre for

efficacy, in comparison to the 5 to 20 gal/acre that can be used for imazapyr to yield equivalent

results. As specified on the product label, glyphosate (as Rodeotm) requires the use of a non-ionic

surfactant for application. These would include surfactants such as R-11 and LI-700, which, as

previously discussed, are one to two orders of magnitude more toxic to rainbow trout than

surfactants such as Hasten or Agridex that can be used with imazapyr (see Table 3-16). Figure 5-1 

depicts the range of potential hazard quotients of four surfactants used with either imazapyr or 

glyphosate under the range of application rates commonly used. This modeling, conducted at an 

assumed water depth of 0.1 m (10 cm or approximately 4 inches) shows how at least one surfactant

used, R-11, has potential to exceed a hazard quotient of 1.0, when equated to the 96-hr LC50

toxicity values developed by Smith et al. (2003) for juvenile rainbow trout. The potential 

exceedance in the sea-surface microlayer on the incoming tide would yield even higher hazard

quotients, if only for a short period until dilution was achieved. However, as demonstrated in 

Figure 5-1, the crop-oil surfactants Agri-Dex and Hasten provide a substantial margin of safety

relative to their application rates to ensure that neither would pose a toxic risk during application, 

even at high application rates above those required for adequate Spartina control with imazapyr. 

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0

0.2

0.4

0.6

0.8

1

1.2

1.4

2.5 5 10 20 40 80

Application Rate (gal/acre)

Hazard Quotient

R-11

LI-700

HASTEN

AGRI-DEX

Figure 5-1: Hazard quotient (HQ) estimation based on rainbow trout LC50 values of 

these surfactants as developed by Smith et al. (2003), where HQ = estimated water 

concentration/LC50. 

5.2 Uncertainties and Data Gaps

Every ecological risk assessment has inherent uncertainties that can only be addressed from

additional studies. The fundamental question in addressing the significance of the uncertainty in

any risk assessment is the degree to which it could qualify the risk conclusions. That is, in this 

report we reviewed the most recent data developed on the toxicity, fate, and degradation of

imazapyr. These results indicated that imazapyr has insignificant toxicity to aquatic and terrestrial 

wildlife, is not environmentally persistent, and does not bioconcentrate or bioaccumulate. While 

the risks to ecological receptors appear extremely low, several data gaps exist. The following 

bullets summarize some of the main data gaps that, if resolved,would eliminate elements of the

uncertainty in this assessment. Uncertainties associated specifically with the manner of

preparation and conclusions of this risk assessment are summarized in Table 5-4.

• Studies pertaining to the effect of imazapyr on aquatic or water-dependent species other 

than fish are limited. No studies examining the toxicity of imazapyr to amphibians and 

reptiles were discovered in our literature review. No studies on the toxicity of imazapyr to 

marine fish typical of those areas where invasive Spartina is distributed in Washington 

State have been conducted. 

• Specific data on the toxicity of imazapyr to sediment-associated organisms typical of northtemperate marine environments is generally lacking and represents a significant data gap. 

• Residues of imazapyr in treated Spartina, and the degradation of the herbicide over time in 

plant tissue were not identified in the literature. Exposure calculations in this assessment

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therefore relied on estimated concentrations in the plant tissue. Empirical residues from 

plants would increase confidence in the exposure and risk estimates. 

• Effects on the micorhizosphere and microflora in a treated estuary have not been explored. 

This subject area has not been investigated thoroughly for any herbicide used in an estuary 

setting to our knowledge. Long term implications of herbicide use on nutrient dynamics

could effect microflora. 

• Effects on non-target salt-marsh plants native to areas Spartina has colonized are poorly 

understood. Limited data on only a few species have been reported. 

• Persistence and stability of imazapyr in dead and decaying Spartina is not known. Can

leachate from decaying vegetation retain herbicidal activity thereby potentially delaying the 

recovery of native salt marsh plants?

• Drift concentrations of imazapyr off-site by treatment method (e.g., backpack, boom 

sprayer, etc.,) have not been quantified. However, worst-case scenario exposure conditions 

in direct application sites did not indicate significant risk. 

• Effects on marine phytoplankton: could herbicide treatments effect nutrient transfer to 

higher trophic levels if phytoplankton are inhibited?

• Effects on sea-surface microlayer associated organisms and microflora in this surface water 

film are not known. 

While the above data gaps represent uncertainty, the existing information on the toxicity and fate

of the compound is substantial and suggests that significant negative impacts would be unlikely in

studies to address the above data gaps—with the possible exceptions of effects on other non-target 

plants and phytoplankton. 

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Table 5-4: Uncertainties associated with ecological risk modeling for imazapyr.

Parameter Source and Results 

Missing 

Information 

(data gaps) 

Information gaps where sources or stressors are not identified or important aspects of the ecology are 

not known can affect risk conclusions. Although it is believed that the important potential sources of 

adverse effects have been addressed, it is possible that there were unmeasured or unconsidered

chemical constituents in the bay that are contributing an unevaluated degree of risk to receptors in 

target areas. 

Conceptual

Model 

If relationships between sources and receptors are missing or incorrectly identified, risks could be

under- or overestimated. To reduce this uncertainty, a conceptual model was developed that identified

all known pathways (both complete and incomplete) and receptor trophic levels. The overall impact of 

this source of uncertainty on risk conclusions is unknown. 

Use of 

Uncertainty

(Safety) Factors

for Calculating 

TRVs 

Uncertainty (safety) factors used to derive TRVs may not accurately reflect site conditions. However, 

the UFs applied were considered realistic based on data from various published studies. Since

published TRVs were not available for all ROIs, UFs were applied. Risk estimates could be under- or 

over-estimated using this approach, as the UFs applied were considered reasonable. 

Laboratory

verses Field 

Populations 

Species used in laboratory toxicity tests are not necessarily subjected to the same degree of nonchemical related stresses as receptors in natural conditions. As such, cumulative effects of multiple

stressors (including chemicals) are not necessarily the same. It is difficult to predict the effect on ERA

results since laboratory versus natural conditions may stress species differently. Due to likely

differences in the health of laboratory populations and those inhabiting target areas, differences in

genetic diversity (hence resistance to stressors), and possible impacts of non-chemical stressors, some

unavoidable uncertainty exists when extrapolating laboratory derived data to field situations. 

Use of 

Representative 

or Surrogate 

Species 

Toxicological studies used species that are related to taxa present in the target areas, but are not

identical. In general, the greater the taxonomic difference, the greater the uncertainty in application of 

laboratory toxicity data to receptors. It is not known whether laboratory test species or receptors in

target areas are the most sensitive to a given chemical constituent. 

Feeding Rates Feeding rates were assumed not to vary with season, breeding condition, or with other local factors. 

Reported feeding rates undoubtedly vary with all of these factors because metabolic needs change as 

does food availability. Where possible, estimates of average feeding rates were derived from studies 

that reported for multiple seasons and areas to compensate for this potential uncertainty. As such, 

while uncertainty is introduced, the effect on ERA conclusions (if any) is not quantifiable. 

5.3 Conclusions and Recommendations 

With current technologies, non-native Spartina eradication cannot be realized in Washington State

without some element of chemical control factored into the WSDA integrated pest management

program. Current mechanical and biological control methods have not been wholly effective, and 

the distribution of Spartina is spreading at a rate of approximately 20 percent per year in some

locations. As Spartina spreads, critical habitat for shorebirds, juvenile fish and shellfish is lost. 

These impacts directly or indirectly impact threatened species such as chinook salmon, and 

commerical enterprises such as shellfish culture. The use of Rodeotm (glyphosate) is the only 

currently approved herbicide for Spartina treatments, but its efficacy is hindered by minimum drytime limits that are not possible under all estuary conditions where Spartina is considered a 

noxious weed. Additional contol means are under investigation, and the use of imazapyr has been

explored in this report as another possible chemical control means. 

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We examined the potential ecological risks from the use of imazapyr in an estuary setting to

control and eventually eradicate non-native Spartina in Washington State. Realistic imazapyr 

exposure scenarios in the estuary settings envisioned for Spartina control did not yield aquatic

concentrations or terrestrial doses that would pose significant risks to aquatic or terrestrial wildlife,

even under the improbable “upper limit” conditions modeled. Only for spill scenarios where it was

assumed that avian and mammalian wildlife could ingest undiluted spray solution was there the 

potential for significant risk identified, but wildlife behavioral mechanisms make this risk scenario 

largely untenable. 

Both imazapyr and glyphosate can be effective at Spartina control and both are essentially safe for

terrestrial and aquatic animals if used in accordance with manufacturers recommendations with

adequate dry-time. However, the use of imazapyr improves the margin of safety relative to 

potentially toxic environmental concentrations by three to four orders of magnitude over the

existing use of glyphosate—depending on which species is considered in the risk assessment. This 

improvement is due to the lower toxicity of imazapyr, lower active ingredient concentration 

needed for Spartina control, lower spray volumes of the herbicide/surfactant solution required for

effective treatment, and the ability to use crop-oil based surfactants which are themselves one to 

two orders of magnitude less toxic than the non-ionic surfactants required for glyphosate use. 

Imazapyr is highly mobile, persistent in soils, and is a broad-spectrum herbicide. Although risks to

animals from imazapyr use are insignificant, its use can cause significant impacts to non-target

vegetation if inappropriately applied. These risks are particularly acute for vascular plants (like

Spartina), although risks to algae appear to be significantly less. In sediment, imazapyr is

significantly less persistent than in soils, but it can still be expected to be detectable for several

weeks after treatment. It should therefore be applied only to target species, avoiding drift or seepage

to non-target species and sediment through observation of weather patterns such as high rains or

wind. Imazapyr should be used primarily in areas where total vegetation control or eradication is 

desired, or in isolated spot applications due to reports of its potential to “leak” out of target plant 

roots into soil that contains non-target plants. Hand spray applications should be used on Spartina

clones and on the periphery of Spartina meadows to minimize spraying of non-target plants and 

poor canopy interception. Broadcast system spraying can be conducted in the central portions of

Spartina meadows with minimal risk of drift and maximum efficacy for Spartina control. 

Photodegradation of imazapyr in water is extremely rapid. In the tidal exhange conditions where 

Spartina is found, dilution is also extremely rapid due to the frequency of tidal exchange. For

example, in Willapa Bay, the primary area where Spartina spp. is distributed and poses the greatest

threat to habitat, there are generally two high and two low tides within a 24-hour period. The

average difference between the high and low tide in Willapa ranges from 8.1 to 10.2 feet, with an

average tidal prism of 4.8 x 108

 cubic yards (cy) and an average tidal flow discharge of 25,000 

cy/second. Potentially toxic concentrations to aquatic animals will not occur under the range of 

application rates considered in this risk assessment and the dilution profiles presented in the 

estuary settings where Spartina occurs. The time of exposure is also reduced because of tidal

exchange rates inherent to Washington’s coastal environments. The overall weight of evidence

from this analysis suggests that imazapyr can be a safe, highly effective treatment for Spartina

control and eradication in an estuary setting, and offers a significantly improved risk scenario over

existing treatment regimes. 

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FINAL 

October 2003 Page 94
Appendix A 

No Action Alternative for Spartina Control 

 
TABLE OF CONTENTS

1.0 INTRODUCTION .................................................................................................................. 1 

2.0 OVERVIEW OF NO ACTION ALTERNATIVE ................................................................. 2 

2.1 Impacts of Spartina .......................................................................................................... 2 

2.2 Description of NAA .......................................................................................................... 3 

3.0 DESCRIPTION OF NO ACTION ALTERNATIVE (NAA) ............................................... 4 

3.1 Spartina Description ......................................................................................................... 4 

4.0 DESCRIPTION OF CURRENT CONTROL METHODOLOGIES AND 

EFFECTIVENESS ..................................................................................................................... 8 

4.1 Physical Removal ............................................................................................................. 8 

4.2 Mechanical ........................................................................................................................ 8 

4.3 Chemical ......................................................................................................................... 10 

4.4 Biological ........................................................................................................................ 11 

5.0 IMPACTS OF CONTROL METHODS ............................................................................... 13 

5.1 Physical Methods ............................................................................................................ 13 

5.2 Mechanical Methods ....................................................................................................... 13 

5.3 Chemical Methods .......................................................................................................... 13 

5.4 Biological Control Methods ........................................................................................... 13 

5.5 Integrated Pest Management Plan .................................................................................. 14 

6.0 DISCUSSION ....................................................................................................................... 18 

7.0 REFERENCES ..................................................................................................................... 22 

LIST OF FIGURES 

Figure A1. Locations of known Spartina spp. infestations in Puget Sound, Hood Canal 

and San Juan Islands (source: WSDA, 2002) .......................................................................... 5 

Figure A2. Locations of known Spartina spp. infestations and treatment sites in 

Willapa Bay (source: WSDA, 2003)........................................................................................ 6 

Figure A3. Locations of known Spartina SPP. infestations in Grays Harbor 

(Source: WSDA 2002) ............................................................................................................. 7

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Appendix A 

No Action Alternative for Spartina Control Page i 
LIST OF TABLES 

Table A-1. Physical Control Methods (source WSDA 1993)........................................................ 8 

Table A-2. Mechanical Control Methods (source WSDA 1993) ................................................ 10 

Table A-3. Chemical (Glyphosate) Control Methods (source WSDA 1993)............................... 11 

Table A-4 . Biological Control Methods (source WSDA 1993) .................................................. 12 

Table A-5a. Potential Physical Habitat Impacts of Different Control Methodologies 

(USFWS 1997)....................................................................................................................... 15 

Table A-5b. Potential Biological Impacts of Control Methodologies (USFWS 1997)............... 16 

Table A-5c. Potential Human Impacts (USFWS 1997)............................................................... 17 

Table A-6. Total Treatment Area and Methodology for Spartina in Washington State 

from 1997 through 2002 (WSDA 2002) ................................................................................ 18 

Table A-6. Total Treatment Area and Methodology for Spartina in Washington State 

from 1997 through 2002 (WSDA 2002), Continued.............................................................. 19 

Table A-7. Potential Efficacy of Treatment Methods on Spartina Infestations in Washington 

State (source: WSDA 1993)................................................................................................... 20 

Table A-8. Summary of the most cost-effective, integrated Spartina Management Practices 

(Norman and Patten, 1997) .................................................................................................... 21

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Appendix A 

No Action Alternative for Spartina Control Page ii 
1.0 INTRODUCTION 

This appendix provides a description of the No Action Alternative (NAA) for smooth cordgrass 

(Spartina spp.) control as practiced by the Washington State Department of Agriculture (WSDA). 

The NAA is described as the program that is presently being employed to eradicate Spartina

species. The Washington State Spartina control program is a cooperative effort between federal,

state, local, tribal and private entities to eradicate this invasive weed. Spartina is rapidly colonizing 

critical mudflat habitat that is used extensively by native shellfish, cultured shellfish, salt marsh 

plants, migratory birds, juvenile fish and other wildlife (WSDA, 2002). The efficacy of current

methods is not as high as desired to control the invasion of this exotic species and Spartina

infestation along Washington State ‘s coastal areas is spreading rapidly. Current control measures

have inherent costs and environmental risks associated with them which additional and/or

alternative methods may alleviate, and thereby facilitate greater efficiency in Spartina control. 

The focus of the NAA description is control of Spartina by herbicides and mechanical and physical

methods. The Ecological Risk Assessment (ERA) presents information for a potential change in

the existing integrated pest management plan (IPMP) for the purpose of providing more effective 

means of Spartina control. The focus of the ERA is to evaluate the risks of using a more effective 

control through changes in the chemical (herbicide) component of the existing program. 

Glyphosate (i.e. Rodeo®) is currently the only chemical registered for aquatic use in the U.S. and 

its use has certain shortcomings. Glyphosate readily binds to sediment particles, which reduces the

translocation (transfer) of chemical down to the roots. Daily inundation by tides cover plants with

mud and dirt, reducing herbicide effectiveness. In addition, application must be timed with tidal 

fluctuations to allow glyphosate to dry in roughly 6 hours. Decreased drying time is directly

related to decreased efficiency or success in eliminating the plant (IVM Technical Bulletin 2003).

Hand spraying with Rodeo requires re-treatment, is poorly adapted to large meadows and the 

logistics of water transport are problematic. The efficacy of broadcast spraying of Rodeo has been 

questionable and the aerial spraying of Rodeo does not work well (Patten 2002). Physical control

methods are labor intensive, expensive, and have small effective areas of control. Mechanical

control is effective if used correctly but is limited by sediment type (soft mud) and by season. 

Changes that have been suggested to improve the present program include a cost-effective 

chemical control and/or a relatively inexpensive large-scale mechanical control (USFWS 1997). 

Patten (2002) suggests that a solution may be a chemical control that is applied in low volumes and 

small concentrations in salt water, that has a minimal non-target impact with no aquatic risk, is able 

to treat large areas with a minimal need for re-treatment and is non-persistent in the water. One 

chemical that may fit these criteria is imazapyr, available in the formulations Arsenaltm and 

Choppertm, amongst others. The ecological hazards and risks from the use of imazapyr are the

primary subject of the Ecological Risk Assessment for which this NAA is appended. 

The objective of this appendix is to present a description of the IPMP in controlling populations of

Spartina spp. in Washington State including effectiveness and potential environmental impacts of

the program. This following sections provide 1) a general background of the Spartina problem in 

Washington State, 2) a description of the present control program, 3) the potential environmental

impacts of the control methods, and 4) the efficacy of these control methodologies. 

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2.0 OVERVIEW OF NO ACTION ALTERNATIVE 

The goal of the NAA is to eradicate Spartina using conventional mechanical, physical, biological

and chemical methods (glyphosate). The problem is that Spartina spreads quickly and is extremely 

difficult to eradicate. The purpose of the NAA program is to control the spread of Spartina using 

methods with minimal environmental impacts and greatest cost effectiveness. 

Prior to the present eradication program, passive management was used to control Spartina spp. 

infestations. Natural processes involving environmental variables, plant genetics and biotic

interactions were thought to regulate distribution and spread of Spartina spp. (WSDA 1993). 

Agencies were to monitor infestations of Spartina spp. and participate in public outreach activities 

under what can be described as a no control alternative. The no control program was characterized 

as not being successful in the eradicating or slowing the spread of Spartina infestations (USFWS 

1997).

The 1995 Legislature designated Washington State Department of Agriculture (WSDA) as the lead 

agency to develop a Statewide Spartina Management Plan. Since that time, WSDA has since

served as the lead state agency for the eradication of Spartina. To accomplish this, six area-wide

Spartina management plans (one for each waterbody covered by a permit) were developed by 

WSDA for North Puget Sound, South Puget Sound, Hood Canal/Central Puget Sound, Grays 

Harbor and Willapa Bay (Figure A1). These programs were developed in conjunction with 

Washington Department of Fish and Wildlife (WDFW), Washington Department of Natural 

Resources (DNR), United States Fish and Wildlife Service (USFWS), Washington State Noxious

Weed Control Board, tribal communities, and private landowners. These management plans

detailed historical information on known infestations, past treatment efforts, and plans for future

control seasons. WSDA consolidated information from the six area-wide management plans into a

single draft document. The Statewide Spartina Management Plan is updated yearly as new control

and survey data/techniques become available. Current methodologies in place to control Spartina

in Washington State include digging, mowing, mowing in combination with herbicide, herbicide 

use only, seedling removal, and various mechanical control methods. 

The NAA presently in operation is referred to as the Integrated Pest Management (IPMP) plan. 

The IPMP is based on the coordinated use of multiple preventative, biological, mechanical, 

physical, and chemical treatments methodologies to control and eradicate the Spartina infestation

in Washington State. 

2.1 IMPACTS OF SPARTINA 

Spartina can alter ecological processes that govern wetland ecosystem function. Infestations can

alter the physical aspect, structure, and spatial configuration of wetlands through dense growth and 

sediment accumulation. The four species now found in Washington State out compete and 

displace beneficial native vegetation and also threaten to severely impact economically important

shellfish cultures. Shellfish are impacted by Spartina encroaching on the available area for oyster 

cultivation and reduction of oyster growth by decreasing the amount of nutrients reaching the 

oyster beds (WSDA 2002). This occurs as the plants trap sediment, which reduces the flow of

water through river channels and changes the elevation of mudflats. Although the amount of 

sediment accumulation is variable from site to site, Spartina's dense root and stems effectively trap 

sediment at higher rates than normal, altering water movement by filling shorelines and river deltas 

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Appendix A 

No Action Alternative for Spartina Control Page 2 
and possibly causing flooding during periods of heavy rain. Spartina spp. destroy important

migratory shorebird and waterfowl habitat by reducing their ability to move, rest and forage on 

open mudflats. Spartina also affects fisheries because fish utilize estuaries as nursing grounds or 

foraging sites. Spartina can also overtake intertidal sea grass beds that provide important habitat

for juvenile fish (USFWS 1997). 

2.2 DESCRIPTION OF NAA 

Spartina poses challenges to implementing control methodologies and is particularly difficult to 

eradicate because it grows in soft mud and has an extensive root network. It also spreads rapidly in

areas that are hard to traverse and apply treatment. These large infestations of Spartina are

difficult to control with present methodologies and no individual method currently being used is

able to effectively control Spartina infestations (USFWS 1997). Currently there are over 15,000 

acres infested in Willapa Bay, over 4,000 acres in Snohomish County and minor infestations in 

eight other counties in western Washington (WSDA 2002). 

The current control program involves active Integrated Pest Management IPM to combat 

infestations of Spartina spp. using mechanical, physical, biological, chemical and/or a combination 

of these methods. The IPMP was chosen as the current program control methodology because it is 

a comprehensive approach that combines a management process with the best components of the 

other methods (WSDA 2002). The current eradication program involves four steps: 

1. Preventing an existing infestation from producing seed; 

2. Treating an existing infestation for several consecutive years using IPM; 

3. After successful eradication is achieved, monitoring the area and removing new 

seedlings to ensure no re-establishment occurs and; and 

4. Continuing to survey shorelines, educate the public and follow-up on possible sightings

of new infestations. 

This is the most promising approach to regionally stabilize and decrease noxious weed 

distributions. The current program involves the deliberate selection, integration, and directed use

of plant population suppression measures on the basis of predicted economic, environmental, and 

sociological consequences. When these measures are successfully applied, plant populations

should be prevented from attaining economically and/or environmentally damaging densities

(USFWS 1997). 

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3.0 DESCRIPTION OF NO ACTION ALTERNATIVE (NAA) 

3.1 SPARTINA DESCRIPTION 

Spartina is commonly known as cordgrass and is an invasive weed that inhabits mudflats, salt 

marshes and estuaries throughout Washington State’s coastal areas. There are now four species of

Spartina found in Washington’s waters including Spartina alterniflora, Spartina patens, Spartina

anglica and a newly discovered species found in the fall of 2001, which is Spartina densiflora. 

Spartina spp. are currently disrupting several native Pacific coastal ecosystems in western

Washington. 

Spartina is a successful invader because of its high rate of spread, its tall and dense canopy that can 

shade out other plants, and its ability to colonize low intertidal regions (Daehler and Strong 1994;

WSDA 1993). The stems are stout and the rhizomes form an extensive root system that is roughly 

five times larger than its aboveground biomass. Once established, Spartina spreads vegetatively,

forming ring-shaped clumps of individual clones. These clones are tall and conspicuous against

open mudflats. New stems grow along the outer edge of the ring, gradually increasing its diameter

with each growing season, while old, dying vegetation can be found in the middle. As clones

spread, they grow into each other, forming a dense Spartina monoculture that overgrows native 

plants (Daehler and Strong 1994). Zipperer (1996) has documented the potential of exotic species 

to change the physical structure and alter the ecological functioning of regional ecosystems.

Spartina follows this pattern well since it out competes and physically displaces native vegetation 

by converting littoral mudflats to salt marshes. This invasion could manifest into large-scale 

ecological changes that threaten to adversely impact fisheries, shellfish beds, waterfowl migrations

and other wildlife dependant on native coastal marshes. 

3.1.1 Origin and Geographic Area 

Spartina alterniflora typifies an invasive species by having a wide tolerance to habitat 

requirements, fast dispersal rate, clonal reproduction (from a single plant spreading asexually) and 

few to no natural predators in its invaded range (Zipperer 1996). Spartina alterniflora was most 

likely introduced to the Washington coast when it was used in the packing of oysters from the East

Coast for shipping during the late 1800s. Spartina alterniflora was also intentionally planted by a 

gun club between 1941 and 1946 to stabilize bank erosion on their property in Padilla Bay. 

Spartina anglica was also intentionally introduced to stabilize dikes and provide forge for cattle in

Port Susan Bay (Fig. A-1). The pathways of introduction for both Spartina patens and the newly

discovered S. densiflora are not currently known. Spartina species have spread throughout coastal

Washington State and presently, ten counties in western Washington have one or more infestations

of Spartina alterniflora, Spartina anglica, Spartina patens or Spartina densiflora (Figures A1 

through A3). These include Clallam, Grays Harbor, Island, Jefferson, King, Kitsap, Pacific, San 

Juan, Skagit and Snohomish counties. Spartina infestations range from one colony in Clallam 

County measuring 50 feet in diameter to more than 7,800 solid acres spread throughout Willapa 

Bay in Pacific County (Figure A2). Spartina infests over 8,000 solid acres and has spread over 

more than 20,000 total acres in Washington State. 

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No Action Alternative for Spartina Control Page 4 
Figure A1. Locations of known Spartina spp. infestations in Puget Sound, Hood Canal and San Juan Islands 

(Source: WSDA 2002) 

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No Action Alternative for Spartina Control Page 5 
Figure A2. Locations of known Spartina spp. infestations and treatment sites in Willapa Bay (Source: WSDA 

2003)

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Figure A3. Locations of known Spartina SPP. infestations in Grays Harbor (Source: WSDA 2002)

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4.0 DESCRIPTION OF CURRENT CONTROL METHODOLOGIES AND 

EFFECTIVENESS 

Current control methods vary in effectiveness, depending on size of infestation, age of plants, 

topography, and proximity to agricultural, aquacultural, or built environments. 

4.1 PHYSICAL REMOVAL 

The Physical Removal Method primarily involves manual labor methods. Current physical control

methods being used include the following: 

• Hand pulling, Digging 

This method is effective for small patches and on young plants. 

• Covering 

This involves layering (covering) Spartina infestations with geo-textile fabric mats that 

inhibit light penetration in order to stop photosynthesis. This method is only used for small

patches of Spartina.

Most physical control methods are labor intensive and time consuming but manual removal can be

effective for removing seedlings and very small infestations or to prevent seed production and 

spread (Table A1). Volunteer organizations and tribal agencies have conducted small-scale

removal projects successfully. 

Table A-1. Physical Control Methods 

(source WSDA 1993)

Physical 

Control 

Method 

Current 

Use 

Most Practical

Applications / Use in present

program Use Constraints 

Hand pulling or

digging 

yes Eradication of small, isolated 

clumps, seedlings, or sparse

infestations / limited use 

Labor-intensive; multiple treatments may be required; plants

must be accessible by foot; pulling limited to seedlings small 

enough to pull 

Covering yes Eradication of small colonies / 

limited use 

Labor-intensive over a large area; wind or waves may dislodge 

covers; monitoring during treatment required; biodegradable 

materials are preferred; non-biodegradable materials will require

removal after use; covers may be buried by sediment

Dewatering/ 

Draining 

no Eradication of medium-sized

colonies, etc

Not practical for large areas or gradual slopes near sea level;

difficulty in obtaining permits; efficacy unknown 

Flooding/ 

inundating

no Eradication of medium- sized 

colonies / etc.

Not practical; shoreline topography must be amenable; difficulty

in obtaining permits

Burning/flaming no Eradication of medium- sized 

colonies

Treatment should be done prior to seed set, during dry weather, 

and when wind blows smoke away from inhabited areas;

smoke might be toxic to workers; flaming is labor-intensive; 

difficulty in obtaining permits 

4.2 MECHANICAL 

This methodology involves the use of machinery to control Spartina from a large area. Machines 

include tools with power sources and range from hand-held brush cutters to amphibious track 

vehicles and barge-mounted dredges. State and federal agencies have aggressively pursued the

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No Action Alternative for Spartina Control Page 8 
treatment of Spartina meadows using mechanical control. The goal of mechanically treating 

Spartina meadows is to reduce current seed production, kill Spartina plants, stress the remaining 

plants sufficiently to reduce seed production in subsequent years, and reduce the amount of

herbicide used in future applications while increasing spray efficacy and reducing cost (USFWS

1997; WSDA 2002). This method is not selective and can damage desirable vegetation and disturb 

soil. This method is normally used only on even terrain with few obstacles. As with manual

removal, control efforts are normally timed to pre-empt seed production. Mechanical control 

efforts are always followed by measures (mechanical or other) to kill shoots from resprouting

plants and seedlings sprouting from the seedbank. Mechanical control methods evaluated in the

EIS (WSDA 1993) include dredging, digging, plowing, rototilling, crushing, mowing, and

harvesting (Table A-2). Current mechanical control methods being used include: 

• Mowing 

Repeated mowing in late spring and summer can prevent seed production, delay colony 

expansion, weaken root vigor and reduce same season re-growth. Mowing can increase

susceptibility to other control methods and can reduce the amount of herbicide needed to 

control Spartina.

• Mechanical sub-soiling and Rototilling 

Mechanical sub-soiling and rototilling involve the mechanical disruption of Spartina, its 

root system, and the supporting substrate by a ripping implement mounted to a amphibious

tracked vehicle (WSDA 1993). This is effective on some upland weeds because it results

in the desiccation of roots. Rototilling can increase susceptibility to other control methods 

and can reduce the amount of herbicide needed to control Spartina. 

• Crushing 

This method destroys the aboveground portion, and can affect the below ground rootmass. 

Crushing is only used in solid meadows of Spartina. Treatment of Spartina clones would 

involve having to transit across open mudflat from one clone to another. Some of the

potential environmental impacts may be minimized since this method is only used on 

meadows and not used on clones. 

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Table A-2. Mechanical Control Methods 

(source WSDA 1993)

Mechanical 

Control 

Method 

Current 

Use Most Practical Applications Use Constraints

Mowing yes Prevention or reduction of seed set; 

delay of expansion of small to large 

monotypic colonies

Multiple treatments required to kill plants; labor

intensive if handheld equipment used; treatment

before seed set 

Digging yes Eradication of small monotypic colonies

where sediment has accreted 

Must be accessible from shore; might not be

possible on soft sediments

Crushing yes Prevention or reduction of seed set; 

reduction of small to large monotypic

colonies. 

Multiple treatments required for eradication; 

crushing by foot should be limited to small clones, 

whereas mechanical crushers can be used for large 

clones; heavy machinery not appropriated for use 

in soft substrates; labor intensive if done by foot;

treatment should be done early in the growing 

season if possible.

Rototilling yes Prevention or reduction of seed set; 

delay of expansion of small to large 

monotypic colonies

Effective during winter but extremely slow; 

expensive to operate machinery

Dredging no Eradication of large monotypic 

infestations where sediment accretion 

has destroyed navigable channels

Must be accessible from water 

Harvesting no Prevention or reduction of seed set Requires market for products; could only be done

where colony spread would not be problematic; 

should be conducted to augment control efforts and

not to maximize yield. 

4.3 CHEMICAL

The Washington State Department of Agriculture (WSDA) has legislative responsibility over the

control of noxious and invasive weeds in the State of Washington. Under this mandate, the agency

is authorized to use herbicides selectively to control invasive weeds. Glyposate (Rodeo®) and 2,4-

dichlorophenoxyacetic acid (2,4-D) were the two chemicals evaluated in the Environmental Impact

Statement (EIS) (WSDA 1993). 2,4-D has not been used since it is not registered for use for 

Spartina control in marine environments in Washington State. Rodeo® has been registered for use

in the marine environment. It is a non-selective herbicide that affects the growth processes in

plants. Glyphosate (the active ingredient) tightly binds to soil particles, reducing its ability to 

contaminate groundwater while microorganisms break it down in water and sediment. Rodeo® is

an effective and relatively non-toxic herbicide that is effective on a wide range of plant species but

it must be used with a nonionic surfactant (R-11® Spreader Acitvator, X-77® Spreader, or LI-700®) 

to help cover the plant and reduce the surface tension of spray droplets (WSDA, 2002). Two

different methodologies are used to apply the herbicide (Table A-3) as summarized below. 

• Herbicide (handheld ground application) 

Ground application of glyphosate from a backpack unit, low pressure or high pressure unit, 

is effective for treating clones and meadows. 

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• Herbicide (broadcast ground application) 

Application by precision sprayer vehicle or airboat is timed to coincide with the maximum 

susceptibility of the plant to increase efficacy (i.e. application at low tide). Broadcast 

application is effective for treating clones and meadows. 

• Herbicide (aerial application) – 

Aerial application by helicopter is timed to coincide with the maximum susceptibility of the 

plant to increase efficacy (i.e., application at low tide). Aerial application is effective at

seed prevention, and in upper elevation Spartina meadows, can reduce plant density. 

Table A-3. Chemical (Glyphosate) Control Methods 

(source WSDA 1993)

Chemical 

Control Most Practical Applications Use Constraints 

Aerial Spray Eradication of large monotypic

infestations where sediment

accretion has destroyed 

navigable channels

Helicopter: quick for large area; spray ball can minimize drift and 

overspray. Boat, Truck, ATV Spraying: more control than helicopter;

reduced possibility of drift overspray; more time and labor than 

helicopter.

Ground 

Application 

Eradication of small monotypic

colonies where sediment has

accreted 

Wicking: may require less herbicide for control; no possibility of 

overspray; may require several passes across plant; labor intensive. 

Back-pack Spray: almost twice as rapid application as wicking; 

easier than wicking; some drift or overspray could occur. 

In several studies, glyphosate applications have had mixed effectiveness (efficacy ranged from 0 to

100 percent ) in controlling Spartina. Short drying times cause the tides to wash glyphosate off the

treated plants and thus limit its efficacy in controlling Spartina. A recent study in Willapa Bay

found no significant reduction in Spartina stem density when applied by helicopter using a 5 

percent solution. However, the same concentration of Rodeo® (5 percent Rodeo®, 95 percent

water/surfactant) applied with a hand-held wand resulted in an 84 percent reduction in Spartina 

stem density (WSDA 2002). Killbride et al., (1995) also indicated that ground rather than aerial

treatments obtained greater control. Ground treatments also increased herbicide contact by

applying higher concentrations of chemical with a brush and by the cleaning action of wiping the

chemical on the plant leaves. Wiping herbicide onto plants both cleans the dirt off plants and 

applies herbicide, but is labor intensive. Ground, compared to aerial applications decreases the 

amount of herbicide drift. 

Applying the herbicide to plants before seeds are produced and when the weed is most susceptible

is crucial to the effectiveness of the treatment. The herbicide must be applied during an outgoing 

tide for maximum drying time. Glyphosate is effective for clone and seedling control if using high 

rate and spray volume. It is also good for high meadows where drying time is greater than 12 hours

but is problematic in intertidal meadows with short dry times (Patten 2002). Decreased drying 

time usually results in decreased efficiency (IVM Technical Bulletin 1987). 

4.4 BIOLOGICAL 

The Biological methods involve the use of biologically based controls, such as pathogens, insects,

livestock, genetic engineering, and competitive plant species to manage infestations of noxious 

species (Table A-4). The purpose of biological control is not to eradicate weeds, but to reduce the 

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infestation and keep them at low, manageable levels. After their introduction, biocontrol agents can 

take 5 to 12 years to become established and increase to numbers large enough to cause damage. 

There is also a long time frame to complete testing and gain regulatory approvals. Once

established, effective biological controls provide an inexpensive, long-term, and non-toxic means

to control weed populations. The planthopper, Prokelisia marginata, was selected as the most 

promising natural enemy because of its known potency against Spartina alterniflora and its narrow 

host range. This insect sucks the sap from the leaf veins of Spartina, depleting its energy supply. 

In addition, P. marginata harms Spartina by inserting its eggs between the leaf layers, scarring the

leaves and causing structural damage to the vascular system. Approximately 200,000 planthoppers

were released in 2001 and observations indicated these insects have reproduced and their offspring 

are currently feeding and developing on the plants (Grevstad 2001). Prokelisia marginata, had an 

unusually devastating effect on Spartina alterniflora from Willapa Bay and damage to the plants is

just beginning to be visible (Grevstad 2001). Released of Prokelisia marginata were made to a

remote population of Spartina anglica in Puget Sound during 2003. Potential insect agents for 

control of S. densiflora and S. patens have not been investigated. 

Direct grazing by domestic livestock could affect the health and vigor of Spartina infestations and 

might be a possible biological control mechanism where the mudflat substrate would support

grazing livestock. 

Table A-4 . Biological Control Methods 

(source WSDA 1993)

Biological 

Control 

Method Most Practical Applications Use Constraints 

Target Insect Planthopper, Prokelisia marginata, holds 

promise for large-scale control of seed 

production in Spartina alterniflora 

Need to release insects at many sites throughout the 

infested area to build up high densities needed to control 

Spartina 

Livestock 

Grazing 

Eradication of small colonies

monospecific stands of Spartina

Soft mud substrate will hinder grazing 

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5.0 IMPACTS OF CONTROL METHODS

5.1 PHYSICAL METHODS 

Environmental impacts associated with physical methods include soil erosion, sediment

mobilization, non-target species mortality, noxious plant dispersal, soil compaction, disruption of 

aquatic food webs, and safety of laborers. The severity of impacts is dependent both on the

specific physical method chosen and size of area treated. Impacts associated with all physical

methods would be expected to increase with the size of area treated. Depending on treatment

method, many impacts can be mitigated by using appropriate procedures and materials and by 

carefully timing treatments (USFWS 1997). 

5.2 MECHANICAL METHODS 

Significant impacts include noxious plant dispersal, soil compaction, sediment mobilization, nontarget plant and animal mortality, increased water turbidity, and disruption of aquatic food webs. 

The extent of these impacts varies with treatment method and size of treatment area. Many 

impacts can be mitigated by using appropriate procedures and materials, and by carefully timing 

treatments. Harvesting does not halt infestation spread and requires a market for plant products,

but could be beneficial as a method of controlling seed production. Digging, rototilling, crushing,

and/or mowing would be applicable in some situations for some species (USFWS 1997). 

5.3 CHEMICAL METHODS 

The potential impacts from use of glyphosate result from the toxicity of the herbicide or from other

factors, such as loss of vegetation, algal blooms, loss of oxygen in water, and erosion and sediment

instability from loss of plant cover. The magnitude of the toxic impacts depends on the 

concentration of glyphosate in the environment, the toxicity of glyphosate, and the extent to which 

humans, wildlife, and nontarget plants could be exposed to glyphosate. Although no significant

impacts from glyphosate are expected for wildlife and humans, there could be toxic effects on

eelgrass and algae. 

Biodegradation and toxicity studies in fish, invertebrates, and mammals of the surfactants approved 

for use in Washington and chemicals similar to the surfactants, indicate that the environmental 

concentrations of surfactants that would result from spraying are not expected to have significant

adverse impacts to human health or the environment. If the herbicide is not applied appropriately, 

it could affect non-target vegetation. However, results of human health risk assessments indicate

that the concentrations of glyphosate to which the public could be exposed are expected to be 

below levels of concern. (USFWS 1997). 

5.4 BIOLOGICAL CONTROL METHODS 

There is an ecological risk involved in introduction of non-indigenous bio-control organisms.

These organisms might cause possible death or injury of native species or agricultural grasses. 

Livestock grazing might negatively impact sediments, soils, water quality, and non-target biota . 

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5.5 INTEGRATED PEST MANAGEMENT PLAN 

Impacts due to implementation of the IPMP control methods can have combined impacts from 

implementation of the individual methods, with some method combinations having additional

synergistic effects. Combination of methods can expose Spartina to multiple stresses, resulting in a 

reduction in environmental impacts and frequency of use of each control methodology (USFWS 

1997) 

Tables A-5a through A5c summarize the impacts of the different control methodologies on 

Willapa Bay (USFWS 1997). These tables are excerpted from the EA conducted by USFWS in

1997 (EA based on the 1993 EIS for western Washington coastal areas). These impacts are

specific to Willapa Bay but they can be generalized and extrapolated to include all of the Spartina

infested areas along the Washington coast. 

The USFWS, (1997) has indicated that Spartina is changing habitat for fish species. Spartina

converts usable intertidal habitat to high meadow that is available to fish only during the highest

tides. There is concern that the Spartina meadows are less complex from an ecological perspective

and will negatively impact fish. There are also concerns that control methods negatively impact

fish. The primary concern is that chemicals used to kill Spartina might impact or compromise fish

health. 

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Table A-5a. Potential Physical Habitat Impacts

of Different Control Methodologies

(USFWS 1997)

Physical 

Issues IPMP Physical/Mechanical Only Chemical Only

Soils and 

Topography

Spartina-facilitated sediment

buildup in scattered clones and 

meadows would be slowed or 

halted in the shortest amount of

time. Low concentrations of 

chemicals would bind to soil and 

then break down.

Spartina-facilitated sediment 

buildup in scattered clones would 

be slowed or halted, but some

meadows would continue to

capture sediment

for many years 

Spartina-facilitated sediment buildup 

in scattered clones and meadows

would be slowed or halted, but over 

a longer period of time than with 

IPMP. Chemicals would bind to soil

and then break down. 

Hydrology Alteration of natural flow patterns 

would be slowed over the next 

few years and reversed in the 

long- run. 

Alteration of natural flow patterns 

would likely be slowed over the 

next few years. Changes in water 

movement due to Spartina would 

continue in areas where control

could not be accomplished. 

Alteration of natural flow patterns 

would be slowed over the next 

decade, and possibly reversed 

eventually. 

Water Quality Potential for short-term herbicide

contamination. Greatest potential

for reducing temperature 

increases and changes in salinity

and oxygen levels

caused by Spartina. 

Greatest localized increases in

suspended sediments. Reduced 

potential for temperature

increases and changes in salinity

and oxygen levels, but to a lesser 

degree than other methods. 

Potential for herbicide 

contamination. Reduced potential

for temperature increases and 

changes in salinity and oxygen 

levels. 

Ambient 

Sound

Short term increased ambient 

noise levels associated with use 

of aircraft, airboats, weed cutters, 

etc.

A higher reliance on large

machinery that could work night 

and day would likely result in

more noise than other methods. 

Even with use of aircraft, this 

method would generate less total 

noise than other methods mainly

because of reduced work

opportunity. Ground-based chemical 

application machinery tends to

generate less noise than 

mechanical methods.

Air Quality Of methods, least potential for air 

pollution from combustion of

fossil fuels due to higher 

efficiency. 

Less potential for herbicide drift

than Chemical Only Method. 

Of methods, highest potential for 

pollution from combustion of

fossil fuels. 

Of methods, there would be a higher 

potential for herbicide spray drift due 

to total reliance on chemical control. 

Pollution from burning fossil fuels 

may be comparable to that of the 

Physical/Mechanical Method.

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Table A-5b. Potential Biological Impacts of Control Methodologies 

(USFWS 1997)

Biological 

Issues PMP Physical/Mechanical Only Chemical Only

Vegetation The greatest acreage of 

native plant habitat would be 

preserved and eventually 

restored. All existing mudflat 

habitat supporting

macroalgae and eelgrass 

would be maintained. Some 

Spartina dominated areas 

would be converted to 

mudflat and some would 

convert to native saltmarsh. 

Existing native saltmarsh 

would be preserved. Impacts 

to vegetation due to Spartina

spread would be minimized. 

Of the methods, this will 

preserve the smallest acreage of

native plant habitat. Mudflats 

supporting eelgrass and 

macroalgae would be 

maintained. Some Spartina 

meadow would likely remain and 

expand on tidelands. Less 

efficient at controlling Spartina 

tidelands would have more 

impacts to adjacent vegetation 

than other methods. 

Mudflat supporting eelgrass and 

macroalgae on the would be 

maintained. Most of the existing 

Spartina meadow would convert 

to native saltmarsh. Seed 

production would likely be 

stopped over time, reducing 

impacts an adjacent areas. 

Spartina Spartina would not increase. 

Seed production could be 

stopped within a short 

timeframe. Spartina 

meadows and clones would 

be mostly eliminated. 

Spartina expansion would be 

slowed. Seed production would 

be stopped. Some Spartina

meadows would likely remain 

after years of control effort. 

Seed production would likely be 

stopped in about a decade. 

Meadows would be eliminated 

and greatly reduced. 

Wildlife Existing mudflat and native 

saltmarsh habitat would be 

maintained. Former areas of 

this habitat, now occupied by 

Spartina would revert mostly 

to native saltmarsh. Wildlife 

use would be sustained. 

Some existing mudflat habitat 

would continue to be lost to 

Spartina. Nearly all Spartina

meadow killed by this method 

would convert to native 

saltmarsh. Of the methods, this 

would preserve the least mudflat 

habitat for migratory bird use. 

Most of the existing mudflat and 

native saltmarsh habitat would 

be maintained. Former areas of 

this habitat, now occupied by 

Spartina would revert mostly to 

native saltmarsh. Wildlife use 

would be sustained. 

Fish More habitat would be 

protected for existing fish 

populations than in other 

methods. 

Some habitat would become 

unusable for fish due to this 

method’s likely inability to fully 

control Spartina. 

Most of the habitat would be 

protected for existing fish 

populations. 

Invertebrat

es

Potential for trampling of 

invertebrates at work sites. 

More habitat for the existing 

invertebrate community 

would be protected than with 

other methods. 

Potential for trampling of 

invertebrates at work sites. 

Where Spartina is not controlled, 

species composition would 

change. 

This action would have the least 

potential for trampling of 

invertebrates at work sites. Most 

of the habitat for the 

existing invertebrate community 

would be protected. 

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Table A-5c. Potential Human Impacts 

(USFWS 1997)

Social 

Issues IPMP Physical/Mechanical Only Chemical Only 

Human 

Health

Health problems associated 

with Spartina pollen 

production and 

Mosquitos that breed in 

Spartina meadows would be 

reduced by Spartina control. 

This method provides the 

greatest opportunity to reduce 

such impacts.

There could be some reduction 

in pollen production and in 

mosquito habitat however, this 

method’s inability to fully control 

Spartina would allow these 

problems to persist to a degree. 

Greater risk of injury to control 

workers. 

Health problems associated with 

Spartina pollen production and 

mosquitos that breed in Spartina 

meadows would be reduced by 

this method, but to a slightly 

lesser extent than IPMP. 

Concerns Concerns about chemical use 

would be reduced to the 

degree that control would not 

depend 

Exclusively on chemical 

methods. Spartina spread 

would be stopped, reducing 

concerns over the loss of 

aquatic resources. 

The absence of chemical use 

would remove concerns about 

chemical risk. Spartina spread 

would be slowed, but not as 

much as with IPMP or chemical 

only. 

Concerns about chemical risk 

would be greatest under this 

method. Concerns about Spartina 

spread would be reduced. 

Recreation Noise from airboats, 

hovercraft, and aircraft. 

Disturbance of waterbirds 

would reduce bird 

Observations. In the long 

term, would be beneficial to 

recreational uses. 

Due to reduced control, some 

recreational opportunities would 

decline. Noise disturbance to 

recreational users would like be 

greater than with other methods. 

Noise would be generated by 

airboats, hovercraft, and aircraft, 

but over shorter periods of time. 

Disturbance of waterbirds would 

reduce bird observations. In the 

long term, would be beneficial to 

recreational uses. 

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6.0 DISCUSSION 

WSDA has been developing regional management plans since 1998. The Statewide Spartina

management plan provides information for the effects of Spartina on the intertidal ecology of these

areas, describe previous control efforts/results, and outline the control strategy for the coming years 

(Table A-6). In 2002, the WSDA Spartina eradication program worked collaboratively with 

partner agencies to continue Spartina control, as outlined in five regional integrated pest 

management plans. The program included hiring, equipping and coordinating workers to treat 

infestations in Clallam, Jefferson, Kitsap and King counties. WSDA also assisted the Swinomish 

and Suquamish tribal communities with control work on their property and worked cooperatively 

with the WDFW and DNR on infestations in Willapa Bay. WSDA worked cooperatively with 

Ecology to develop a NPDES permit for aquatic noxious weed control, providing NPDES 

coverage to numerous federal, state and local governmental agencies, and private entities for

herbicide applications to both marine and freshwater (WSDA, 2002). 

Table A-6. Total Treatment Area and Methodology for Spartina 

in Washington State from 1997 through 2002 (WSDA 2002) 

County Spartina Present in 2002

Spartina Treated, 

1997 - 2002 2002 Treatment Methods

Pacific (Willapa Bay) Over 6,800 solid acres 

spread over > 15,000 acres 

‘97 – approx. 742 solid acres 

‘98 – approx. 450 solid acres 

‘99 – approx. 600 solid acres 

‘00 – approx. 800 solid acres 

‘01 – approx. 900 solid acres 

‘02 – approx. 1804 solid acres 

Mow/herbicide, herbicide, 

seedling removal, various 

mechanical control 

(rototilling, sub-soiling, etc).

Grays Harbor Scattered clones and 

seedlings 

0.25 acres in size 

‘97 – all treated 

‘98 - all treated 

‘99 – all treated 

‘00 – all treated 

‘01 – all treated 

‘02 – all treated 

Herbicide, seedling removal, 

mow 

Snohomish Approx. 350 solid acres 

spread over > 4,500 acres 

‘97 – approx. 89 solid acres 

‘98 – approx. 126 solid acres 

‘99 – approx. 90 solid acres 

‘00 – approx. 158 solid acres 

‘01 – approx. 75 solid acres 

‘02 – approx. 238 solid acres 

Mow/herbicide, herbicide, 

seedling removal, dig, 

mechanically crush, mow 

Island Approx. 350 solid acres 

spread over >1,000 acres 

‘97 – approx. 250 solid acres 

‘98 – approx. 160 solid acres 

‘99 – approx. 155 solid acres 

‘00 – approx. 130 solid acres 

‘01 – approx. 72 solid acres 

‘02 – approx. 180 solid acres 

Mow/herbicide, herbicide, 

seedling removal, 

mechanically crush, mow 

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Table A-6. Total Treatment Area and Methodology for Spartina 

in Washington State from 1997 through 2002 (WSDA 2002), Continued 

County Spartina Present in 2002

Spartina Treated, 

1997 - 2002 2002 Treatment Methods

Skagit Approx. 40 solid acres 

spread over > 2,000 acres 

‘97 – approx. 91 solid acres 

‘98 – approx. 57 solid acres 

‘99 – all treated 

‘00 – approx. 60 solid acres 

‘01 – approx. 33 solid acres 

‘02 – approx. 37 solid acres 

Mow/herbicide, herbicide, 

seedling removal, dig, mow 

Clallam 1 infestation < 0.001 acres 

in size 

‘97 – treated twice 

‘98 – treated three times 

‘99 – treated twice 

‘00 – treated three times 

‘01 – treated four times 

‘02 – treated four times 

Dig 

Jefferson 14 infestations – approx. 

0.01 solid acres total 

‘97 - all treated 

‘98 - all treated twice 

‘99 – all treated twice 

‘00 – all treated twice 

‘01 – all treated three times 

‘02 – all treated three times 

Mow, mow/herbicide, dig, 

seedling removal 

Kitsap 8 infestations - approx. 1 

solid acre total 

‘97 - all but 2 tribal sites 

‘98 - all treated 

‘99 – all treated twice 

‘00 – all treated 

‘01 – all treated 

‘02 – all treated twice 

Mow mow/herbicide, dig, 

seedling removal 

King 2 infestations – single 

clones and a few seedlings 

‘97 – monitored 

‘98 – all treated 

‘99 – all treated 

‘00 – all treated twice 

’01 – all treated twice 

‘02 – all treated twice 

Dig 

San Juan Re-growth found at one 

site. 2 other sites clean for 

four consecutive years 

‘97 - all treated 

‘98 - all treated 

‘99 – monitored 

‘00 – all treated 

’01 – all treated 

‘02 – all treated 

Survey, dig 

The potential efficacy of the control methods is outlined in Table A-7. Mowing and herbicide

application seem to have the highest efficacy for containment and reduction of Spartina. 

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Table A-7. Potential Efficacy of Treatment Methods on Spartina

Infestations in Washington State (source: WSDA 1993) 

Potential Efficacy/

Infestation Threat Objective Management Method Small Area 

Large 

Area Plant Growth Stage 

Herbicide Low to High Low to 

High 

Actively growing, 

especially pre-flowering 

Cutting/mowing Medium to

High 

Medium to 

High 

Pre-flowering or early 

flowering, every month 

Mechanical crushing Low to High Low to 

High 

Pre-flowering or early 

flowering, May through 

August 

Established 

Invader Core 

Containment, 

Reduction 

Covering Medium Low All stages during growing 

season 

Herbicide Low to High Low to 

High 

Seedlings, pre-flowering 

Hand pulling Medium to 

High 

Medium to 

High 

Seedlings to small plants 

Established 

Invader Outliers 

Containment or 

Control 

Covering High Medium to

High 

All stages during growing 

season, treatments 

repeated every month 

Herbicide Low to High Low to 

High 

Seedling, pre-flowering 

Hand pulling/digging Medium to 

High 

Medium Seedling to small plants 

Covering High Medium to

High 

All stages during growing 

season, treatments 

repeated every month 

New Invader Eradication, 

Reduction, 

Containment, 

Control 

Mechanical crushing Low to High Low to 

High 

Pre-flowering or early 

flowering, May through 

August 

Education/awareness Medium to 

High 

Medium to 

High 

All Infestations Prevention 

(applies to all 

infestations) Surveys Medium to

High 

Medium to 

High 

Patten (1997) has indicated that mowing followed by herbicide application provides the highest 

efficacy but at an increased cost (Table A-8). Aerial application is more cost effective and covers 

wider areas but the efficacy is questionable. 

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Table A-8. Summary of the most cost-effective, 

integrated Spartina Management Practices (Norman and Patten, 1997) 

Method % Kill (Efficacy) Acreage $/Acre

Mow only 95 low 312 

Mow + Rodeo 98 low 431 

Rodeo hand wipe 91 low 310 

Rodeo hand spray 81 low 585 

Rodeo aerial spray ? high 165 

WSDA has demonstrated that Spartina eradication is feasible. The current strategies for control in

each region are founded upon the 1998 plans. Many of these plans are proving successful, 

especially in North Puget Sound where the strategy has resulted in a 27% decline in the overall size

of the infestation (WSDA, 2001a; WSDA, 2002). The overall Puget Sound infestation was

reduced by 10% from 1997 to 1999, Spartina establishment in Grays Harbor has been prevented, 

and populations of Spartina at select sites in Willapa Bay have been eradicated (WSDA, 2001b;

WSDA, 2002). This is the positive aspect concerning the present program, but the Spartina

infestation continues to grow. At current levels, it could take decades to eradicate Spartina in 

Puget Sound and it may never be eradicated in Willapa Bay (WSDA, 2002). 

Methodologies and equipment for eradicating Spartina have evolved over time with treatment

efforts. The agencies now use airboats to transport equipment and personnel, large-scale

amphibious mowing machines to stop seed production, small tracked vehicles to shred and rip 

apart isolated infestations, high pressure spray systems to treat large clones and fringes of 

meadows, and volunteers, landowners and students to dig seedlings (WSDA 2002). WSDA

(2002), concluded that, large-scale mechanical eradication of Spartina is not feasible at this time. 

A potential alternative at this time, might be an effective chemical control. The chemical control 

solution would need to be a cost-effective chemical that can be applied in low volumes and small

concentrations in salt water and would integrate minimal non-target impact with acceptable aquatic 

risk and non-persistent in the water. This would allow for large areas to be treated with a minimal

need for re-treatment. 

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7.0 REFERENCES 

Daehler, C.C. and D.R. Strong. 1994. Variable reproductive output among clones of Spartina 

alterniflora (Poacea) invading San Francisco Bay, California: the influence of herbivory, 

pollination, and establishment Site. American Journal of Botany 81(3):307-313. 

Grevstad, F. 2001. Spartina Biocontrol—Background and Progress. University of Washington, 

Olympic Natural Resources Center. Website: 

http://www.willapabay.org/~coastal/nospartina/biocontrol/biocontrol_overview2001.pdf

IVM Technical Bulletin, 1998. Smooth Cordgrass (Spartina). In Integrated Pest Management

Practitioner's Association: Bio-Integral Resource Center (BIRC)'s IPMP Access: Noxious

Weed Integrated Vegetation Management Guide: IVM Technical Bulletin. Website: 

http://www.efn.org/~ipmpa/Noxspart.html. Visited: 6/5/03. 

Killbride, K.M, F.L. Paveglio, and C.E. Grue. 1995. Control of smooth cordgrass with Rodeo in a 

southwestern Washington estuary. Wildlife Society Bulletin 23(3): 520-524. 

Major III, W. and C.E. Grue. 1997. Control of Spartina alterniflora in Willapa Bay, Washington: 

efficacy of mechanical and chemical control techniques and their off target impacts on 

eelgrass (Z. japonica). Second International Spartina Conf. Proc. 1997: 76-81. 

Norman, M. and K. Patten. 1997. Cost-efficacy of integrated Spartina control practices in Willapa 

Bay, Washington. 1997. Second International Spartina Conference Proceedings 1997: 89-

92. 

Patten, K. 2002. Imazapyr- efficacy, toxicity, costs and registration. Presentation at Spartina

Conference, May 13-14. Olympia, WA. Website: 

http://www.willapabay.org/~coastal/nospartina/conferences/conf2002/conference_may2002.htm Visited: 

6/5/03. 

U.S. Department of Agriculture (USDA). 1995. Imazapyr: Pesticide Fact Sheet. Prepared by 

Ventures, Inc. November 1995. Website: http://infoventures.com/e-hlth/pestcide/imazapyr.html 

Visited: 6/5/03. 

U.S. Fish and Wildlife Service (USFWS). 1997. Environmental Assessment for Control of Smooth 

Cordgrass (Spartina Altierniflora) on Willapa National Wildlife Refuge. U.S. Fish and 

Wildlife Service. Washington. 

Washington State Departments of Agriculture, (WSDA). 1993. Noxious emergent plant 

management: final environmental impact statement. State of Washington. Olympia. 204 pp 

+ app.

Washington State Department of Agriculture (WSDA). 2000, Spartina and Purple Loosestrife 

Report to the Legislature Eradication and Control Programs (as required by RCW 

17.26.015). 44 pp. 

J:\Divisions\Plant Protection\Spartina\HabitatEcoRiskAssessment\Appendix A Rpt.doc

Appendix A 

No Action Alternative for Spartina Control Page 22
Washington State Department of Agriculture (WSDA). 2001a. 2001 Spartina Management Plan 

for North Puget Sound. Prepared by WSDA in cooperation with the North Puget Sound 

Spartina Task Force. 

Washington State Department of Agriculture (WSDA). 2001b. 2001 Spartina Management Plan 

for Willapa Bay. Prepared by WSDA in cooperation with the Willapa Bay Spartina Task 

Force. 

Washington State Department of Agriculture (WSDA). 2002. Report to the Legislature. State of 

Washington. Olympia. 

Zipperer, V.T. 1996. Ecological Effects of the Introduced Cordgrass, Spartina alterniflora, on the 

Benthic Community Structure of Willapa Bay, Washington. Masters of Science Thesis. 

University of Washington. 

J:\Divisions\Plant Protection\Spartina\HabitatEcoRiskAssessment\Appendix A Rpt.doc

Appendix A 

No Action Alternative for Spartina Control Page 23
Appendix B 

Exposure Calculation Worksheets 

 
APPENDIX B:

Exposure Calculation Worksheets

Dermal 

Absorbed Dose

Terrestrial Receptor AR mg/cm2 Body Weight SA Upper Amnt Typical Amnt Upper D

Deer Mouse 0.0168 0.021 90.11 1.14 0.76 54.0

Norway Rat 0.0168 0.300 507.52 6.39 4.26 21.3

Cottontail Rabbit 0.0168 1.286 1307.16 16.47 10.98 12.8

Red Fox M 0.0168 5.250 3261.57 41.10 27.40 7.83 

Red Fox F 0.0168 4.130 2790.55 35.16 23.44 8.51 

Marsh Wren 0.0168 0.011 60.06 0.76 0.50 67.2

Bobwhite Quail 0.0168 0.190 377.15 4.75 3.17 25.0

Greater Scaup M 0.0168 0.860 1006.35 12.68 8.45 14.7

Greater Scaup F 0.0168 0.770 936.57 11.80 7.87 15.3

Mallard Duck M 0.0168 1.225 1266.52 15.96 10.64 13.0

Mallard Duck F 0.0168 1.043 1140.80 14.37 9.58 13.7

W = body weight (kg); SA = exposed surface areaEstimated Absorbed Dose = Dabs = Amnt/WTypical Amount deposited on the animal (Amnt) = 0.5 x SA x AR Upper Amount Deposited On the Animal (Amnt) = 0.75 x SA x AR Application Rate AR= 0.0168 mg/cm2

Exposed Surface Area SA= 1110(W)0.65

Amt Deposited on Animal (mg) Dose A

Page 1 of 1 
Ecological 

Receptor

Acute DietaryUpper Exposure1 

Acute DietaryTypical Exposure1 

Chronic Dietary Upper Exposure2

Chronic Dietary Typical Exposure2

Dermal/ ContactUpper Exposure3

Dermal/ ContactTypical Exposure3

Drinking Water (Spill) Upper Exposure4

Drinking Water (Spill) Typical Exposure4

Deer Mouse 0.0084214 0.000524 0.014946429 0.007842857 0.02703254 0.0180217 1.19773333 0.5988666Norway Rat 0.000524 0.000172 0.00093 0.000488 0.01065789 0.0071053 0.395252 0.19762Cottontail Rabbit 0.0027504 0.0009028 0.012203538 0.006403577 0.13516268 0.0901085 0.34926128 0.1746306Red Fox M 0.0007226 0.0002372 0.005130057 0.002691901 0.05328943 0.0355263 0.3018288 0.150914Red Fox F 0.0007232 0.0002374 0.00513414 0.002691901 0.03201837 0.0213456 0.3088586 0.154429Marsh Wren 0.0552852 0.018147 0.066133333 0.034702222 0.01956941 0.0130463 7.07675744 3.5383787Bobwhite Quail 0.0115083 0.0037775 0.013766447 0.007223684 0.02128389 0.0141893 2.80587225 1.4029361Scaup M 0.0152326 0.0008703 0.005406977 0.002837209 0.16816289 0.1121086 0.96362661 0.4818133Scaup F 0.017013 0.000972 0.006038961 0.003168831 0.06252731 0.0416849 0.99503125 0.4975156Mallard Duck M 0.1002551 0.0057281 0.035586735 0.018673469 0.03686032 0.0245735 0.8552047 0.4276023Mallard Duck F 0.1177493 0.0067277 0.0417965 0.021931927 0.03831438 0.0255429 0.65962899 0.3298141: acute dietary reference NOEL = 5,000 mg/kg-bw for mouse, rat, rabbit, and fox; 1149 mg/kg for mallard and scaup; and 674 mg2: chronic reference NOEL dose for mouse and rate = 1,000 mg/kg; for rabbits = 400 mg/kg; for dogs = 250 mg/kg; for avian spec3: dermal/contact NOEL dose for mouse & rat= 2,000 mg/kg; 400 mg/kg for rabbit. No data on fox or avian species, so assumed 4: same reference doses as acute oral 
C upper in mg/L C lower in mg/L A W Upper Typical

Deer Mouse 3.4 0.1 0.007 0.021 1.133 0.033

Norway Rat 3.4 0.1 0.033 0.3 0.374 0.011

Cottontail Rabbit 3.4 0.1 0.125 1.286 0.330 0.010

Red Fox M 3.4 0.1 0.441 5.25 0.286 0.008

Red Fox F 3.4 0.1 0.355 4.13 0.292 0.009

Marsh Wren 3.4 0.1 0.003 0.0113 0.903 0.027

Bobwhite Quail 3.4 0.1 0.02 0.19 0.358 0.011

Scaup M 3.4 0.1 0.053 0.86 0.210 0.006

Scaup F 3.4 0.1 0.049 0.77 0.216 0.006

Mallard Duck M 3.4 0.1 0.067 1.225 0.186 0.005

Mallard Duck F 3.4 0.1 0.044 1.043 0.143 0.004

Dose is calculated as (C x A)/WA = water consumed per day (in L); and W = average body weight (in kg) C = concentration of neat solution in spill (mg/L) based on application rates of 10 gal/acre (upper), and 20 gal/acre (typical)

[note: lower application volumes result in higher neat concentrations of herbicide, hence higher concentr

Ecological 

Receptor

Dose Ingested from 
C upper in mg/L C lower in mg/L A W Upper Typical

Deer Mouse 17966 8983 0.007 0.021 5988.7 2994.3

Norway Rat 17966 8983 0.033 0.3 1976.3 988.1

Cottontail Rabbit 17966 8983 0.125 1.286 1746.3 873.2

Red Fox M 17966 8983 0.441 5.25 1509.1 754.6

Red Fox F 17966 8983 0.355 4.13 1544.3 772.1

Marsh Wren 17966 8983 0.003 0.0113 4769.7 2384.9

Bobwhite Quail 17966 8983 0.02 0.19 1891.2 945.6

Scaup M 17966 8983 0.053 0.86 1107.2 553.6

Scaup F 17966 8983 0.049 0.77 1143.3 571.6

Mallard Duck M 17966 8983 0.067 1.225 982.6 491.3

Mallard Duck F 17966 8983 0.044 1.043 757.9 379.0

Dose is calculated as (C x A)/WA = water consumed per day (in L); and W = average body weight (in kg) C = concentration of neat solution in spill (mg/L) based on application rates of 10 gal/acre (upper), and 20 gal/acre (typical)

[note: lower application volumes result in higher neat concentrations of herbicide, hence higher concen

Dose Ingested from Spill Solution (mg/kg-bw)

Terrestrial 

Ecological 

Receptor
Chronic Dietary Dose

Terrestrial Receptor Ctwa upper Ctwa typical

food intake/day 

(kg/1000)) Prop (W)

Upper

 Limit Dose 

Deer Mouse 46.5 24.4 9 0.75 21 14.95 Norway Rat 46.5 24.4 15 0.4 300 0.93 Cottontail Rabbit 46.5 24.4 180 0.75 1286 4.88 Red Fox M 46.5 24.4 362 0.4 5250 1.28 Red Fox F 46.5 24.4 285 0.4 4130 1.28 Marsh Wren 46.5 24.4 8 0.4 11.25 13.23 Bobwhite Quail 46.5 24.4 15 0.75 190 2.75 Greater Scaup M 46.5 24.4 50 0.4 860 1.08 Greater Scaup F 46.5 24.4 50 0.4 770 1.21 Mallard Duck M 46.5 24.4 250 0.75 1225 7.12 Mallard Duck F 46.5 24.4 250 0.75 1043 8.36 Time-weighted average concentration on Vegetation (Ctwa) = Co(1-e-kt)/(kT) (note: units in mg/kg)food intake per day is in kg/1000, from Wildlife exposure handbookW = body weight in kg/1000 Chronic Dietary Dose Absorbed (CD) = (Ctwa)(food intake/day)(Prop)/W 
Acute Dietary Dose

Terrestrial Receptor C upper C typical A Prop W

Upper Dose 

(mg/kg-bw)

Typi

(mg

Deer Mouse 131 43 9 0.75 21 42.11 1

Norway Rat 131 43 15 0.4 300 2.62 

Cottontail Rabbit 131 43 180 0.75 1286 13.75 

Red Fox M 131 43 362 0.4 5250 3.61 

Red Fox F 131 43 285 0.4 4130 3.62 

Marsh Wren 131 43 8 0.4 11.25 37.26 1

Bobwhite Quail 131 43 15 0.75 190 7.76 

Greater Scaup M 131 43 50 0.4 860 3.05 

Greater Scaup F 131 43 50 0.4 770 3.40 

Mallard Duck M 131 43 250 0.75 1225 20.05 

Mallard Duck F 131 43 250 0.75 1043 23.55 

Dose= AxC(Prop)/WA = daily food intake in gramsProp is unitless estimate of percent of total diet assumed from contaminated vegetationC = residue concentration in food (mg/kg)W = body weight in grams
Appendix C 

Avian Species in Willapa Bay

 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name

LOONS Spring Summer F

Red-throated Loon GAVIA STELLATA c-

Pacific Loon GAVIA PACIFICA crCommon Loon GAVIA IMMER cr

GREBES Spring Summer F

Pied-billed Grebe* PODILYMBUS PODICEPS uuHorned Grebe PODICEPS AURITUS crRed-necked Grebe PODICEPS GRISEGENA r -

Western Grebe AECHMOPHORUS OCCIDENTALIS au

FULMARS, PETRELS AND SHEARWATERS Spring Summer F

Northern Fulmar FULMARUS GLACIALIS -r

Pink-footed Shearwater PUFFINUS CREATOPUS --

Sooty Shearwater PUFFINUS GRISEUS uc

Short-tailed Shearwater PUFFINUS TENUIROSTRIS --

STORM PETRELS Spring Summer F

Fork-tailed Storm-Petrel OCEANODROMA FURCATA --

Leach's Storm-Petrel* OCEANODROMA LEUCORHOA --

PELICANS AND CORMORANTS Spring Summer F

Brown Pelican PELECANUS OCCIDENTALIS oc

Double-crested Cormorant* PHALACROCORAX AURITUS ccBrandt's Cormorant* PHALACROCORAX PENICILLATUS cc

BITTERNS, HERONS AND EGRETS Spring Summer F

American Bittern* BOTAURUS LENTIGINOSUS ou

Great Blue Heron* ARDEA HERODIAS cc

Great Egret ARDEA ALBA o-Cattle Egret BUBULCUS IBIS --

Green Heron BUTORIDES VIRESCENS rr

Abundance by s

Page 1 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by s

WATERFOWL Spring Summer F

Tundra Swan CYGNUS COLUMBIANUS - - 

Trumpeter Swan CYGNUS BUCCINATOR - - 

Greater White-fronted Goose ANSER ALBIFRONS o-

Snow Goose CHEN CAERULESCENS o-Ross' Goose CHEN ROSSII r-

Emperor Goose CHEN CANAGICA r-

Brant BRANTA BERNICLA aoCanada Goose* BRANTA CANADENSIS acWood Duck* AIX SPONSA uuGreen-winged Teal ANAS CRECCA crMallard* ANAS PLATYRHYNCHOS ccNorthern Pintail ANAS ACUTA urBlue-winged Teal ANAS DISCORS urCinnamon Teal* ANAS CYANOPTERA uuNorthern Shoveler ANAS CLYPEATA urGadwall ANAS STREPERA urEurasian Wigeon ANAS PENELOPE - -

American Wigeon ANAS AMERICANA cr

Canvasback AYTHYA VALISINERIA u-Ring-necked Duck AYTHYA COLLARIS u-Tufted Duck AYTHYA FULIGULA --

Greater Scaup AYTHYA MARILA u-

Lesser Scaup AYTHYA AFFINIS c-Harlequin Duck HISTRIONICUS HISTRIONICUS r-

Oldsquaw CLANGULA HYEMALIS o-

Black Scoter MELANITTA NIGRA u-Surf Scoter MELANITTA PERSPICILLATA c o White-winged Scoter MELANITTA FUSCA coCommon Goldeneye BUCEPHALA CLANGULA u-Barrow's Goldeneye BUCEPHALA ISLANDICA r-

Page 2 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sBufflehead BUCEPHALA ALBEOLA c-Hooded Merganser* LOPHODYTES CUCULLATUS uoCommon Merganser* MERGUS MERGANSER cuRed-breasted Merganser r c cRuddy Duck OXYURA JAMAICENSIS o - 

VULTURES Spring Summer F

Turkey Vulture CATHARTES AURA uu

OSPREY, KITES, EAGLES AND HAWKS Spring Summer F

Osprey* PANDION HALIAETUS uu

White-tailed Kite ELANUS LEUCURUS ou

Bald Eagle* HALIAEETUS LEUCOCEPHALUS uuNorthern Harrier* CIRCUS CYANEUS ccSharp-shinned Hawk ACCIPITER STRIATUS urCooper's Hawk ACCIPITER COOPERII urNorthern Goshawk ACCIPITER GENTILIS r-

Red-tailed Hawk* BUTEO JAMAICENSIS cc

Rough-legged Hawk BUTEO LAGOPUS u-

FALCONS Spring Summer F

American Kestrel FALCO SPARVERIUS urMerlin FALCO COLUMBARIUS u-Peregrine Falcon FALCO PEREGRINUS u-Gyrfalcon FALCO RUSTICOLUS --

GALLINACEOUS BIRDS Spring Summer F

Ring-necked Pheasant* PHASIANUS COLCHICUS uu

Blue Grouse* DENDRAGAPUS OBSCURUS uuRuffed Grouse* BONASA UMBELLUS uuWild Turkey MELEAGRIS GALLOPAVO rr

Northern Bobwhite* COLINUS VIRGINIANUS uu

Page 3 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by s

RAILS Spring Summer F

Virginia Rail* RALLUS LIMICOLA uu

Sora PORZANA CAROLINA r-

American Coot FULICA AMERICANA u-

PLOVERS Spring Summer F

Black-bellied Plover PLUVIALIS SQUATAROLA cuAmerican Golden Plover PLUVIALIS DOMINICA rr

Snowy Plover* CHARADRIUS ALEXANDRINUS uu

Semipalmated Plover CHARADRIUS SEMIPALMATUS ccKilldeer* CHARADRIUS VOCIFERUS uu

OYSTERCATCHERS Spring Summer F

American Oystercatcher* HAEMATOPUS PALLIATUS uu

SHOREBIRDS Spring Summer F

Greater Yellowlegs TRINGA MELANOLEUCA cu

Lesser Yellowlegs TRINGA FLAVIPES --

Willet CATOPTROPHORUS SEMIPALMATUS r -

Wandering Tattler HETEROSCELUS INCANUS uoSpotted Sandpiper ACTITIS MACULARIA uoWhimbrel NUMENIUS PHAEOPUS coLong-billed Curlew NUMENIUS AMERICANUS u-Bar-tailed Godwit LIMOSA LAPPONICA --

Marbled Godwit LIMOSA FEDOA uo

Ruddy Turnstone ARENARIA INTERPRES coBlack Turnstone ARENARIA MELANOCEPHALA uuSurfbird APHRIZA VIRGATA crRed Knot CALIDRIS CANUTUS c-Sanderling CALIDRIS ALBA acSemipalmated Sandpiper CALIDRIS PUSILLA or

Page 4 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sWestern Sandpiper CALIDRIS MAURI aaLeast Sandpiper CALIDRIS MINUTILLA ccPectoral Sandpiper CALIDRIS MELANOTOS --

Sharp-tailed Sandpiper CALIDRIS ACUMINATA r-

Dunlin CALIDRIS ALPINA auStilt Sandpiper CALIDRIS HIMANTOPUS --

Ruff PHILOMACHUS PUGNAX --

Short-billed Dowitcher LIMNODROMUS GRISEUS aaLong-billed Dowitcher LIMNODROMUS SCOLOPACEUS ur

SNIPE Spring Summer F

Common Snipe GALLINAGO GALLINAGO cr

PHALAROPES Spring Summer F

Wilson's Phalarope PHALAROPUS TRICOLOR --

Red-necked Phalarope PHALAROPUS LOBATUS uo

Red Phalarope PHALAROPUS FULICARIA rr

JAEGERS Spring Summer F

Parasitic Jaeger STERCORARIUS PARASITICUS rr

GULLS AND TERNS Spring Summer F

Bonaparte's Gull LARUS PHILADELPHIA cu

Heermann's Gull LARUS HEERMANNI oc

Mew Gull LARUS CANUS crRing-billed Gull LARUS DELAWARENSIS cuCalifornia Gull LARUS CALIFORNICUS cuHerring Gull LARUS ARGENTATUS --

Thayer's Gull LARUS THAYERI --

Western Gull* LARUS OCCIDENTALIS ccGlaucous-winged Gull* LARUS GLAUCESCENS ccBlack-legged Kittiwake RISSA TRIDACTYLA ur

Page 5 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sSabine's Gull XEMA SABINI rr

Caspian Tern* STERNA CASPIA cc

Common Tern STERNA HIRUNDO urArctic Tern STERNA PARADISAEA r-

SEABIRDS Spring Summer F

Common Murre URIA AALGE uc

Pigeon Guillemot* CEPPHUS COLUMBA ccMarbled Murrelet* u u uAncient Murrelet SYNTHLIBORAMPHUS ANTIQUUS --

Cassin's Auklet PTYCHORAMPHUS ALEUTICUS --

Rhinoceros Auklet CERORHINCA MONOCERATA ouTufted Puffin FRATERCULA CIRRHATA ouHorned Puffin FRATERCULA CORNICULATA --

DOVES Spring Summer F

Rock Dove* COLUMBA LIVIA uu

Band-tailed Pigeon* COLUMBA FASCIATA ccMourning Dove ZENAIDA MACROURA rr

OWLS Spring Summer F

Barn Owl* TYTO ALBA uu

Western Screech-Owl* OTUS KENNICOTTII uuGreat Horned Owl* BUBO VIRGINIANUS uuSnowy Owl NYCTEA SCANDIACA --

Northern Pygmy-Owl* GLAUCIDIUM GNOMA uu

Burrowing Owl - r Barred Owl* STRIX VARIA uuLong-eared Owl ASIO OTUS r-

Short-eared Owl ASIO FLAMMEUS uo

Northern Saw-whet Owl* AEGOLIUS ACADICUS uu

Page 6 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by s

GOATSUCKERS Spring Summer F

Common Nighthawk* CHORDEILES MINOR ru

SWIFTS Spring Summer F

Vaux's Swift* CHAETURA VAUXI cc

HUMMINGBIRDS Spring Summer F

Anna's Hummingbird CALYPTE ANNA --

Rufous Hummingbird* SELASPHORUS RUFUS aa

KINGFISHERS Spring Summer F

Belted Kingfisher* CERYLE ALCYON uu

WOODPECKERS Spring Summer F

Red-breasted Sapsucker SPHYRAPICUS RUBER u-

Downy Woodpecker* PICOIDES PUBESCENS uu

Hairy Woodpecker* PICOIDES VILLOSUS uuNorthern Flicker* COLAPTES AURATUS ccPileated Woodpecker* DRYOCOPUS PILEATUS uu

FLYCATCHERS Spring Summer F

Olive-sided Flycatcher* CONTOPUS BOREALIS ccWestern Wood-Pewee* CONTOPUS SORDIDULUS uuWillow Flycatcher* EMPIDONAX TRAILLII uuPacific-slope Flycatcher* EMPIDONAX DIFFICILIS ccWestern Kingbird TYRANNUS VERTICALIS r-

LARKS Spring Summer F

Horned Lark* EREMOPHILA ALPESTRIS uu

SWALLOWS Spring Summer F

Purple Martin* o r -

Page 7 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sTree Swallow* TACHYCINETA BICOLOR ccViolet-green Swallow* TACHYCINETA THALASSINA ccNorthern Rough-winged SwallowSTELGIDOPTERYX SERRIPENNIS uu

Cliff Swallow* HIRUNDO PYRRHONOTA cc

Barn Swallow* HIRUNDO RUSTICA ca

JAYS, MAGPIES AND CROWS Spring Summer F

Gray Jay PERISOREUS CANADENSIS oo

Steller's Jay* CYANOCITTA STELLERI uu

Scrub Jay APHELOCOMA COERULESCENS o o 

American Crow* CORVUS BRACHYRHYNCHOS cc

Common Raven* CORVUS CORAX uu

CHICKADEES AND TITMICE Spring Summer F

Black-capped Chickadee* POECILE ATRICAPILLUS cc

Chestnut-backed Chickadee* POECILE RUFESCENS cc

BUSHTITS Spring Summer F

Bushtit* PSALTRIPARUS MINIMUS or

NUTHATCHES Spring Summer F

Red-breasted Nuthatch SITTA CANADENSIS ur

CREEPERS Spring Summer F

Brown Creeper* CERTHIA AMERICANA u u 

WRENS Spring Summer F

Bewick's Wren* THRYOMANES BEWICKII uu

Winter Wren* TROGLODYTES TROGLODYTES cc

Marsh Wren* CISTOTHORUS PALUSTRIS cc

Page 8 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by s

KINGLETS, BLUEBIRDS AND THRUSHES Spring Summer F

Golden-crowned Kinglet* REGULUS SATRAPA ccRuby-crowned Kinglet REGULUS CALENDULA crWestern Bluebird SIALIA MEXICANA r-

Mountain Bluebird SIALIA CURRUCOIDES r-

Townsend's Solitaire MYADESTES TOWNSENDI orSwainson's Thrush* CATHARUS USTULATUS ccHermit Thrush CATHARUS GUTTATUS u-American Robin* TURDUS MIGRATORIUS ccVaried Thrush* IXOREUS NAEVIUS cu

WAGTAILS AND PIPITS Spring Summer F

American Pipit ANTHUS RUBESCENS --

WAXWINGS Spring Summer F

Cedar Waxwing* BOMBYCILLA CEDRORUM uc

SHRIKES Spring Summer F

Northern Shrike LANIUS EXCUBITOR o-

STARLINGS AND MYNAS Spring Summer F

European Starling* STURNUS VULGARIS cc

VIREOS Spring Summer F

Solitary Vireo* VIREO SOLITARIUS r-

Hutton's Vireo* VIREO HUTTONI uuWarbling Vireo* VIREO GILVUS uu

WARBLERS Spring Summer F

Orange-crowned Warbler* VERMIVORA CELATA ccYellow Warbler* DENDROICA PETECHIA uuYellow-rumped Warbler* DENDROICA CORONATA cu

Page 9 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sBlack-throated Gray Warbler* DENDROICA NIGRESCENS ccTownsend Warbler DENDROICA TOWNSENDI c -Hermit Warbler DENDROICA OCCIDENTALIS r r 

Palm Warbler DENDROICA PALMARUM --

MacGillivray's Warbler OPORORNIS TOLMIEI r r 

Common Yellowthroat* GEOTHLYPIS TRICHAS cc

Wilson's Warbler* WILSONIA PUSILLA cc

TANAGERS Spring Summer F

Western Tanager* PIRANGA LUDOVICIANA uu

GROSBEAKS AND BUNTINGS Spring Summer F

Black-headed Grosbeak* PHEUCTICUS MELANOCEPHALUS uu

TOWHEES AND SPARROWS Spring Summer F

Rufous-sided Towhee* PIPILO ERYTHROPHTHALMUS uu

Chipping Sparrow SPIZELLA PASSERINA r-

Savannah Sparrow* PASSERCULUS SANDWICHENSIS cc

Fox Sparrow PASSERELLA ILIACA u-Song Sparrow* MELOSPIZA MELODIA ccLincoln's Sparrow MELOSPIZA LINCOLNII r-

White-throated Sparrow ZONOTRICHIA ALBICOLLIS oo

Golden-crowned Sparrow ZONOTRICHIA ATRICAPILLA c-White-crowned Sparrow* ZONOTRICHIA LEUCOPHRYS ccDark-eyed Junco* JUNCO HYEMALIS ccLapland Longspur CALCARIUS LAPPONICUS r-

Snow Bunting PLECTROPHENAX NIVALIS - -

BLACKBIRDS, MEADOWLARKS AND ORIOLES Spring Summer F

Red-winged Blackbird* AGELAIUS PHOENICEUS cc

Western Meadowlark* STURNELLA NEGLECTA uu

Yellow-headed Blackbird XANTHOCEPHALUS XANTHOCEPHALUS r-

Page 10 of 11 
APPENDIX C:

Avian Species in Willapa Bay

Species Scientific Name Abundance by sBrewer's Blackbird* EUPHAGUS CYANOCEPHALUS ccBrown-headed Cowbird* MOLOTHRUS ATER ccBullock's Oriole* ICTERUS BULLOCKII o o 

FINCHES Spring Summer F

Purple Finch* CARPODACUS PURPUREUS ccHouse Finch* CARPODACUS MEXICANUS ccRed Crossbill* LOXIA CURVIROSTRA ucCommon Redpoll CARDUELIS FLAMMEA --

Pine Siskin* CARDUELIS PINUS co

American Goldfinch* CARDUELIS TRISTIS uc

WEAVER FINCHES Spring Summer F

House Sparrow* PASSER DOMESTICUS c c 

Notes: a = abundant, c = common, u = uncommon, o = occasional, r = rare, * = known to nest in the area"

Page 11 of 11 
Appendix D 

Terrestrial and Amphibious Wildlife 

in the Wallapa Bay Watershed 

 
APPENDIX D:

Terrestrial and Amphibious Wildlife in the Wallapa Bay Watershed

MARSUPIALS

Virginia opossum DIDELPHIS VIRGINIANA

INSECTIVORES

Vagrant Shrew SOREX VAGRANS

Dusky Shrew SOREX MONTICOLUS

Marsh Shrew NEOSOREX PALUSTRIS

Trowbridge's Shrew SOREX TROWBRIDGII

Shrew-Mole NEUROTRICHUS GIBBSII

Townsend's Mole SCAPANUS TOWNSENDII

Coast Mole SCAPANUS ORARIUS

BATS

Little Brown Myotis MYOTIS LUCIFUGUS

Yuma Myotis MYOTIS YUMANENSIS

Long-eared Myotis MYOTIS EVOTIS

Long-legged Myotis MYOTIS VOLANS

California Myotis MYOTIS CALIFORNICUS

Silver-haired Bat LASIONYCTERIS NOCTIVAGANS

Big Brown Bat EPTESICUS FUSCUS

Hoary Bat LASIURUS CINEREUS

RABBITS AND HARES

Snowshoe Hare LEPUS AMERICANUS

RODENTS

Mountain Beaver APLODONTIA RUFA

Townsend's Chipmunk TAMIAS TOWNSENDII

Douglas Squirrel TAMIASCIURUS DOUGLASII

Northern Flying Squirrel GLAUCOMYS SABRINUS

Beaver CASTOR CANADENSIS

Deer Mouse PEROMYSCUS MANICULATUS

Forest Deer Mouse PEROMYSCUS MANICULATUS GRACILIS

Bushy-tailed Woodrat NEOTOMA CINEREA

Southern Red-backed Vole CLETHRIONOMYS GAPPERI

Townsend's Vole MICROTUS TOWNSENDII

Long-tailed Vole MICROTUS LONGICAUDUS

Oregon Vole MICROTUS OREGONI

Muskrat ONDATRA ZIBETHICUS

Norway Rat RATTUS NORVEGICUS

Pacific Jumping Mouse ZAPUS TRINOTATUS

Porcupine ERETHIZON DORSATUM

Nutria MYOCASTOR COYPUS

CARNIVORES

Coyote CANIS LATRANS

Black Bear URSUS AMERICANUS

Raccoon PROCYON LOTOR

Pine Marten MARTES AMERICANA

Long-tailed Weasel MUSTELA FRENATA

Page 1 of 3 Appendix D
APPENDIX D:

Terrestrial and Amphibious Wildlife in the Wallapa Bay Watershed

Mink MUSTELA VISON

Striped Skunk MEPHITIS MEPHITIS

River Otter LUTRA CANADENSIS

Bobcat LYNX RUFUS

Cougar FELIS CONCOLOR

WHALES, DOLPHINS, AND PORPOISES

Gray Whale ESCHRICHTIUS ROBUSTUS

Harbor Porpoise PHOCOENA PHOCOENA

SEALS AND SEA LIONS

Stellar Sea Lion EUMETOPIAS JUBATUS

California Sea Lion ZALOPHUS CALIFORNIANUS

Harbor Seal PHOCA VITULINA

Northern Fur Seal CALLORHINUS URSINUS

DEER

Roosevelt Elk CERVUS ELAPHUS ROOSEVELTI

Black-tailed Deer ODOCOILEUS HEMIONUS COLUMBIANUS

Columbian White-tailed Deer ODOCOILEUS VIRGINIANUS LEUCURUS

AMPHIBIANS

BULLFROG RANA CATESBEIANA

CASCADE TORRENT SALAMANDER RHYACOTRITON CASCADAE

CASCADES FROG RANA CASCADAE

COLUMBIA SPOTTED FROG RANA LUTEIVENTRIS

COLUMBIA TORRENT SALAMANDER RHYACOTRITON KEZERI

COPE'S GIANT SALAMANDER DICAMPTODON COPEI

DUNN'S SALAMANDER PLETHODON DUNNI

ENSATINA ENSATINA ESCHSCHOLTZII

GREAT BASIN SPADEFOOT SPEA INTERMONTANA

GREEN FROG RANA CLAMITANS

LARCH MOUNTAIN SALAMANDER PLETHODON LARSELLI

LONG-TOED SALAMANDER AMBYSTOMA MACRODACTYLUM

NORTHERN LEOPARD FROG RANA PIPIENS

NORTHWESTERN SALAMANDER AMBYSTOMA GRACILE

OLYMPIC TORRENT SALAMANDER RHYACOTRITON OLYMPICUS

OREGON SPOTTED FROG RANA PRETIOSA

PACIFIC GIANT SALAMANDER DICAMPTODON TENEBROSUS

PACIFIC TREEFROG HYLA REGILLA

RED-LEGGED FROG RANA AURORA

ROCKY MOUNTAIN TAILED FROG ASCAPHUS MONTANUS

ROUGHSKIN NEWT TARICHA GRANULOSA

TAILED FROG ASCAPHUS TRUEI

TIGER SALAMANDER AMBYSTOMA TIGRINUM

VAN DYKE'S SALAMANDER PLETHODON VANDYKEI

WESTERN REDBACK SALAMANDER PLETHODON VEHICULUM

WESTERN TOAD BUFO BOREAS

WOOD FROG RANA SYLVATICA

WOODHOUSE'S TOAD BUFO WOODHOUSII

Page 2 of 3 Appendix D
APPENDIX D:

Terrestrial and Amphibious Wildlife in the Wallapa Bay Watershed

REPTILES

CALIFORNIA MOUNTAIN KINGSNAKE LAMPROPELTIS ZONATA

COMMON GARTER SNAKE THAMNOPHIS SIRTALIS

GOPHER SNAKE PITUOPHIS CATENIFER

GREEN SEA TURTLE CHELONIA MYDAS

LEATHERBACK SEA TURTLE DERMOCHELYS CORIACEA

LOGGERHEAD SEA TURTLE CARETTA CARETTA

NIGHT SNAKE HYPSIGLENA TORQUATA

NORTHERN ALLIGATOR LIZARD ELGARIA COERULEA

NORTHWESTERN GARTER SNAKE THAMNOPHIS ORDINOIDES

PACIFIC GOPHER SNAKE PITUOPHIS CATENIFER CATENIFER

PAINTED TURTLE CHRYSEMYS PICTA

POND SLIDER PSEUDEMYS SCRIPTA

RACER COLUBER CONSTRICTOR

RINGNECK SNAKE DIADOPHIS PUNCTATUS

RUBBER BOA CHARINA BOTTAE

SAGEBRUSH LIZARD SCELOPORUS GRACIOSUS

SHARPTAIL SNAKE CONTIA TENUIS

SHORT-HORNED LIZARD PHRYNOSOMA DOUGLASSI

SIDE-BLOTCHED LIZARD UTA STANSBURIANA

SNAPPING TURTLE CHELYDRA SERPENTINA

SOUTHERN ALLIGATOR LIZARD ELGARIA MULTICARINATA

STRIPED WHIPSNAKE MASTICOPHIS TAENIATUS

WESTERN FENCE LIZARD SCELOPORUS OCCIDENTALIS

WESTERN POND TURTLE CLEMMYS MARMORATA

WESTERN RATTLESNAKE CROTALUS VIRIDIS

WESTERN SKINK EUMECES SKILTONIANUS

WESTERN TERRESTRIAL GARTER SNAKE THAMNOPHIS ELEGANS

Page 3 of 3 Appendix D
Appendix E 

Threatened and Endangered Species and 

Species of Concern within Washington State, 

and the 

Potential for Imazapyr Exposure from Spartina Treatment 

 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Cascades Frog Rana cascadae Amphibian FCo SM N Mountain meadows, streams, ponds, and lakes 

water and vegetation around it.

Columbia Spotted Frog Rana luteiventris Amphibian FCo SC P Mountainous areas near cold streams and lakes

overland in spring and summer.

Columbia Torrent Salamander

Rhyacotriton kezeri Amphibian FCo SC N Live on the edges of cold, clear mountain stream

or in waterfall spray zones. Sometimes found aseason.

Larch Mountain Salamander

Plethodon larselli Amphibian FCo SS N Prefers lava talus slopes and outcrops in dense 

(30-1,189 m).

Northern Leopard Frog Rana pipiens Amphibian FCo SE N From freshwater sites with profuse vegetation to 

fields; from desert to mountain meadow. Not fo

Olympic Torrent Salamander

Rhyacotriton olympicus

Amphibian FCo SM N Live on the edges of cold, clear mountain stream

or in waterfall spray zones. Sometimes found aseason.

Oregon Spotted Frog Rana pretiosa Amphibian FC SE N Mountainous areas near cold streams and lakes

overland in spring and summer.

Red-legged Frog Rana aurora Amphibian FCo none P Usually found near ponds or other permanent w

Also likes damp woods.

Tailed Frog Ascaphus truei Amphibian FCo SM N Usually clear, cold swift-flowing mountain stream

water in damp forests or in more open areas in 

Van Dyke's Salamander Plethodon vandykei Amphibian FCo SC N Streams, mountain seeps, waterfall splash zone

5,000' (457-1,524 m).

Western Toad Bufo boreas Amphibian FCo SC P Near springs, streams, meadows, woodlands o

Beller's Ground Beetle Agonum belleri Beetle FCo SC N A terrestrial species, not found in estuary habitaHatch's Click Beetle Eanus hatchii Beetle FCo SC N A terrestrial species, not found in estuary habitaAleutian Canada Goose Branta canadensis 

leucopareia

Bird FCo ST P An uncommon visitor to coastal 

Page 1 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

American Peregrine Falcon

Falco peregrinus anatum

Bird FCo SS P Open country, especially along rivers; also near 

cities. Migrates chiefly along coasts. Breeds frosouth locally through mountainous West, and spnorth to British Columbia and Massachusetts. Several Subspecies also represented by the Wthese include the Peale's Peregrine.

Bald Eagle Haliaeetus 

leucocephalus

Bird FT ST Y Common resident to Puget Sound and Coastal 

Black Tern Childonias niger Bird FCo SM P Freshwater marshes and marshy lakes in summ

and in winter. No sightings recorded in Willapa

Brown Pelican Pelecanus 

occidentalis

Bird FE SE P Resident of Pacific Coast from southern Califor

northward as far as southern British Columbia a

Atlantic Coast from North Carolina south to Ven

Burrowing Owl Athene cunicularia Bird FCo SC N Plains, deserts, fields, and airportsCassin's Auklet Ptychoramphus 

aleuticus

Bird FCo SC N Open ocean. Nests on sea cliffs and isolated he

Ferruginous Hawk Buteo regalis Bird FCo ST N Prairies, brushy open country, badlands.Harlequin Duck Histrionicus 

histrionicus

Bird FCo none P Swift-moving streams in summer; rocky, wave-l

winter.

Loggerhead Shrike Lanius ludovicianus Bird FCo SC P Grasslands, orchards, and open areas with sca

woodlands; deserts in the West.

Marbled Murrelet Brachyramphus 

marmoratus

Bird FT ST Y Breeds in coastal rain forests; inshore waters at 

Northern Goshawk Accipiter gentilis Bird FCo SC P Breeds in coniferous forests; winters in farmland

country. Rarely recorded from Willapa.

Olive-sided Flycatcher Contopus borealis Bird FCo none* Y Boreal spruce and fir forests, usually near open

Page 2 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Oregon Vesper Sparrow Pooecetes 

gramineus affinis

Bird FCo SC P Prefers relatively dry and sparsely vegetated ar

used for song perches.

Sage-grouse Centrocercus 

urophasianus

Bird FC ST N A rare shrub-steppe species of the Columbia Pl

disturbance factors have disrupted mature shru

Sharp-tailed Grouse Tympanuchus 

phasianellus

Bird FCo ST N Found in the North Central Cascades at altitude

Short-tailed Albatross Phoebastria albatrus Bird FE SC N An open ocean piscivore. Does not frequent co

Coast. An extremely rare bird.

Slender-billed Whitebreasted Nuthatch Sitta carolinensis aculeata Bird FCo SC Y Large trees in deciduous and mixed forests.Snowy Plover Charadrius 

alexandrinus

Bird FT SE Y Sandy coastal beaches and shore of salt ponds 

Washington to Baha, California

Spotted Owl Strix occidentalis Bird FT SE P Wet, coniferous, old-growth forests; innermost t

Willapa Bay coast retain some adjacent old growsupportable for spotted owl

Streaked Horned Lark Eremophila alpestris 

strigata

Bird FC SC P Grasslands and lowland prairies.

Tufted Puffin Fratercula cirrhata Bird FCo SC P Nests on vertical sea cliffs, in colonies or singly

Willow Flycatcher Empidonax traillii Bird FCo none* P Dense streamside thickets, swampy thickets, w

Yellow-billed Cuckoo Coccyzus 

americanus

Bird FC SC $B,RI P Moist thickets, willows, overgrown pastures, and 

Island Marble Euchloe ausonides 

insulanus

Butterfly FCo SC P Grasslands and Garry oak woodlands.

Makah (Queen Charlotte) Copper

Lycaena mariposa charlottensis

Butterfly FCo SC P Sedge dominated fens (prairie terrain) containin

coastal rainforests of the Olympic Peninsula.

Mardon Skipper Polites mardon Butterfly FC SE P Fescue-dominated native prairie grasslands.Oregon Silverspot Butterfly

Speyeria zerene hippolyta

Butterfly FT SE P Coastal meadows in salt spray zone. Spartina 

habitats where its forage, Viola adunca, might o

Page 3 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Taylor's (Whulge) Checkerspot

Euphydras editha taylori

Butterfly FC SC P Native grasslands, prairies, and oak woodlands 

Sound basin, and the Willamette Valley.

Valley Silverspot Speyeria zerene 

bremnerii

Butterfly FCo SC P Moist Garry Oak ecosystems.

Bull Trout (Coastal/Puget Sound)

Salvelinus confluentus

Fish FT SC Y Coastal and mountain streams; deep pool of lar

rarely brackish and salt water.

Bull Trout Salvelinus 

confluentus

Fish FT SC Y Coastal and mountain streams; deep pool of lar

rarely brackish and salt water.

Bull Trout (Columbia Basin)

Salvelinus confluentus

Fish FT SC Y Coastal and mountain streams; deep pool of lar

rarely brackish and salt water.

Chinook Salmon (Upper Columbia SP)

Oncorhynchus tshawytscha

Fish FE SC N Coastal streams and rivers, intertidal areas , an

Chinook Salmon (Snake R. Fall)

Oncorhynchus tshawytscha

Fish FT SC Y Coastal streams and rivers, intertidal areas , an

Chinook Salmon (Puget Sound)

Oncorhynchus tshawytscha

Fish FT SC Y Coastal streams and rivers, intertidal areas , an

Chinook Salmon (Lower Columbia)

Oncorhynchus tshawytscha

Fish FT SC Y Coastal streams and rivers, intertidal areas , an

Chinook Salmon (Snake R. SP/SU)

Oncorhynchus tshawytscha

Fish FT SC N Coastal streams and rivers, intertidal areas , an

Chum Salmon (Hood Canal SU)

Oncorhynchus keta Fish FT SC Y Coastal waters; enters streams to spawn. Natu

the Columbia drainage and naturally spawned sHood Canal and its tributaries in Olympic Peninand Dungeness Bay are classified as threatene

Chum Salmon (Lower Columbia)

Oncorhynchus keta Fish FT SC Y Coastal streams and rivers, intertidal areas , an

Coastal Cutthroat Oncorhynchus clarki 

clarki

Fish FCo none Y Small, cool headwater streams and larger rivers

form in intertidal areas and inshore marine wate

Page 4 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Packific Hake (C. Puget Sound)

Merluccius productus

Fish FC SC Y Coastal Waters as juveniles: Open waters as ad

Pacific Herring 

(Discovery Bay)

Clupea pallasi Fish FC SC P Frequently offshore, but usually inshore in harb

spawning.

Pacific Herring (Cherry Point)

Clupea pallasi Fish FC SC P Frequently offshore, but usually inshore in harb

spawning.

Pacific Lamprey Lampetra tridentata Fish FCo none P Migrate as adults to ocean. Widespread and oft

shallow waters to 820 ft.

River Lamprey Lampetra ayresi Fish FCo SC P Migrate as adults to ocean. Widespread and oft

shallow waters to 820 ft.

Rock Bass Ambloplites rupestris Fish none none N Vegetated and brushy stream margins of lakes. 

rocky streams

Salish Sucker Catostromus SP. Fish none SM N Pools and runs of small cool headwaters and crSand Roller Percopsis 

transmontana

Fish none SM N Quiet backwater and pool margins of small to la

Sockeye Salmon (Snake R.)

Oncorhynchus nerka Fish FE SC N Streams and rivers; lakes; intertidal areas and o

Sockeye Salmon (Ozette Lake)

Oncorhynchus nerka Fish FT SC N Streams and rivers; lakes; intertidal areas and o

Steelhead (Upper Columbia)

Oncorhynchus mykiss

Fish FE SC P Clear, cool streams and rivers; lakes; intertidal a

Steelhead (Snake River) Oncorhynchus 

mykiss

Fish FT SC P Clear, cool streams and rivers; lakes; intertidal a

Steelhead (Middle Columbia)

Oncorhynchus mykiss

Fish FT SC P Clear, cool streams and rivers; lakes; intertidal a

Steelhead (Lower Columbia)

Oncorhynchus mykiss

Fish FT SC Y Clear, cool streams and rivers; lakes; intertidal a

Westslope Cutthroat Oncorhynchus clarki 

lewisi

Fish FCo none N Small, cool streams and rivers; lakes

Page 5 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Black Right Whale Balaena glacialis Mammal FE SE N A marine pelagic species.Blue Whale Balaenoptera 

musculus

Mammal FE SE N A marine pelagic species.

California Bighorn Sheep Ovis canadensis 

Californiana

Mammal FCo none N Semi-open, precipitous terrain with rocky slopes

from alpine meadow to hot desert.

Cathlamet Pocket Gopher

Thomomys mazama louiei

Mammal FC SC N Prairies to mountain meadows. Known only from 

County. It may now be extinct. 

Columbian White-tailed Deer

Odocoileus virginianus leucurus

Mammal FE SE P Coastal and inland floodplains, in woodlands an

Willapa Bay may be suitable for this species, if range.

Destruction Island Shrew Sorex trowbridgii 

destrructioni

Mammal FCo none N Numerous habitats; most common in moist field

woods.

Fin Whale Balaenoptera 

physalus

Mammal FE SE N A marine pelagic species.

Fisher Martes pennanti Mammal FCo SE N Not coastal--a species of mature, dense foreste

Fringed Myotis Myotis thysanodes Mammal FCo SM N Oak, pinyon, and juniper forests; desert scrub. R

buildings, and other protected locations

Gray Wolf Canis lupus Mammal FT SE N Open tundra and forests

Grizzly Bear Ursus arctos Mammal FT SE N Only an occasional visitor to Washington wilder

Cascades; not coastal

Humpback Whale Megaptera 

novaeangliae

Mammal FE SE N A marine pelagic species.

Kincaid's Meadow Vole Microtus 

pennsylvanicus 

Mammal FCo SM P Lush, grassy fields; also marshes, swamps, wo

mountaintops.

Least Chipmunk Tamias minimus Mammal none none* P Pastures, piney woods, rocky cliffs, and sagebr

open coniferous forests.

Long-eared Myotis Myotis evotis Mammal FCo SM Y Variety of habitats, from sage to high-altitude co

in forested regions. Sometimes roosts in buildin

Page 6 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Long-legged Myotis Myotis volans Mammal FCo SM Y Mainly coniferous forests. In summer, roosts in Lynx Lynx canadensis Mammal FT ST N Deep, coniferous forest interspersed with rocky 

thickets

Mazama (Western) Pocket Gopher

Thomomys mazama Mammal FC SC N Prairies to mountain meadows

Olympic Pocket Gopher Thomomys mazama 

melanops

Mammal FC SC N Prairies to mountain meadows

Pacific Townsend's Bigeared Bat Coryhorhinus townsendii Mammal FCo SC P occur in a variety of habitats from desert shrub 

forests at a wide range of elevations

Pallid Townsend's Bigeared Bat Coryhorhinus Townsendii Mammal FCo SC P occur in a variety of habitats from desert shrub 

forests at a wide range of elevations

Preble's Shrew Sorex preblei Mammal FCo SM P Open areas, woodlands, forestsPygmy Rabbit Brachylagus 

idahoensis

Mammal FE SE N It is patchily distributed in the sagebrush-domin

Sea Otter Enhydra lutris Mammal FCo SE N Coastal waters within a mile (1.5 km) of shore; e

kelp beds and abundant shellfish

Sei Whale Balaenoptera 

borealis

Mammal FE SE N A marine pelagic species.

Shelton Pocket Gopher Thomomys mazama 

couchi

Mammal FC SC P Native grasslands, prairies and oak woodlands 

Small-footed Myotis Myotis ciliolabrum Mammal FCo SM P Usually lives in arid, rocky habitats including forSperm Whale Physeter 

macrocephalus

Mammal FE SE N A marine pelagic species.

Steller Sea Lion Eumetopias jubatus Mammal FT ST N Stellers are found throughout the North Pacific 

California. Unlike California sea lions, Stellers teout in unpopulated areas.

Townsend's Big-eared Bat

Coryhorhinus tonsendii

Mammal FCo SC P Western deserts and dry pine forests.

Page 7 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Washington Ground Squirrel

Spermophilus washingtoni

Mammal FC SC P Sagebrush and grassland habitats in Southeast

of Oregon in Gilliam, Morrow, and Umatilla Cou

Western Gray Squirrel Sciurus griseus Mammal FCo ST P Woodlands and coniferous forests, particularly QWolverine Gulo gulo Mammal FCo SC N A wide range of habitats with circumpolar distrib

montane coniferous habitats.

Woodland Caribou Rangifer tarandus Mammal FE SE N Usually found in small herds in boreal forests fro

Newfoundland

Yelm Pocket Gopher Thomomys mazama 

yelmensis

Mammal FC SC P Native grasslands, prairies and oak woodlands 

Yuma Myotis Myotis yumanensis Mammal FCo none* P Yuma myotis occur in a variety of habitats inclu

and deserts, and forests. 

California Floater Anodonta 

californiensis

Mollusk FCo SC N Shallow areas of clean, clear lakes, ponds and 

elevations and a soft, silty substrate to burrow i

Giant Columbia Spire Snail

Fluminicola columbiana

Mollusk FCo SC N Relatively pristine colder, clear streams of the S

with a high dissolved oxygen content.

Newcomb's Littorine Snail

Algamorda subrotundata

45 FCo SC P Rocky shores in the upper intertidal zone of ma

Fender's Soliperlan Stonefly

Soliperla fenderi Other insect FCo none N Fender's soliperlan stonefly nymphs are exclusi

headwaters of small streams. As adults, stoneflthe shores of the streams.

Lynn's Clubtail Gomphus lynnae Other insect FCo none N Clear streams that have relatively open canopie

Columbia clubtail have only been confirmed aloCounty, Washington.

Green Sea Turtle Chelonia mydas Reptile FT ST N Open-ocean species; no breeding colonies in WLeatherback Sea Turtle Dermochelys 

coreacea

Reptile FE SE N Open-ocean species; no breeding colonies in W

Loggerhead Sea Turtle Caretta caretta Reptile FT ST N Open-ocean species; no breeding colonies in W

Page 8 of 9 
APPENDIX E:

Threatened and Endangered Species and Species of Concern within Wahsington State,

and the Potential for Imazapyr Exposure from Spartina Treatment

Common Name Scientific Name

Animal 

Type

Federal 

Status

State 

Status

Potential 

Estuary 

Presence? Habitat and Ran

Sagebrush Lizard Sceloporus 

graciosus

Reptile FCo SC N Primarily areas of sagebrush and gravelly soils 

from shelter such as stony piles, crevices, animnot supported in estuarine conditions.

Western Pond Turtle Clemmys Marmorata Reptile FCo SE P Ponds and small lakes with abundant vegetatio

moving streams, reservoirs, and occasionally in 

Note: Fco = federal species of concern; FT = federally listed as threatened; FE = federally endangered; FC = Federal Candidate SpeciesSE = State endangered; ST = state threatened; SC = state species of concern© 2003Washington Department of Fish and Wildlife

Page 9 of 9