HANDBOOK OFCHEMICAL RISK ASSESSMENT Health Hazards to Humans, Plants, and Animals ( VOLUME 1 ) - PART 5 ppt

CHAPTER 21Mirex Fish and wildlife resources associated with approximately 51 million ha 125 million acres in the southeastern United States, and with the Great Lakes, especially Lake Ont

CHAPTER 21

Mirex

Fish and wildlife resources associated with approximately 51 million ha (125 million acres)

in the southeastern United States, and with the Great Lakes, especially Lake Ontario, have beennegatively affected by intensive or widespread use of mirex, a chlorinated hydrocarbon compound(Waters et al 1977; Bell et al 1978; Kaiser 1978; National Academy of Sciences [NAS] 1978;Lowe 1982; Eisler 1985; Hill and Dent 1985; Sergeant et al 1993; Blus 1995; U.S Public HealthService [USPHS] 1995) Contamination of the Southeast and of Lake Ontario by mirex probablyoccurred between 1959 and 1978 During that period, mirex was used as a pesticide to controlthe red imported fire ant (Solenopsis invicta) and the black imported fire ant (Solenopsis richteri),which infested large portions of Alabama, Arkansas, Florida, Georgia, Louisiana, Mississippi,North Carolina, South Carolina, and Texas Under the trade name of Dechlorane, mirex was used

as a fire retardant in electronic components, fabrics, and plastics; effluents from manufacturingprocesses resulted in the pollution of Lake Ontario Regulatory agencies, environmentalists, andthe general public became concerned as evidence accumulated demonstrating that mirex wasassociated with high death rates, numerous birth defects, and tumors, and that it disruptedmetabolism in laboratory mammals, birds, and aquatic biota Mirex also tends to bioaccumulateand to biomagnify at all trophic levels of food chains Field studies corroborated the laboratoryfindings and showed that mirex appeared to be one of the most stable and persistent organochlo-rine compounds known, being resistant to chemical, photolytic, microbial, metabolic, and thermaldegradation processes Upon degradation, a series of potentially hazardous metabolites areformed, although it is generally acknowledged that the fate and effects of the degradation productsare not fully understood Mirex was also detected in human milk and adipose tissues at lowconcentrations, the levels related to the degree of environmental contamination In 1978, the U.S.Environmental Protection Agency banned all uses of mirex It is probable that mirex and itsmetabolites will continue to remain available to living organisms in this country for at least

12 years, although some estimates range as high as 600 years

Mirex is a white, odorless, free-flowing, crystalline, nonflammable, polycyclic compound posed entirely of carbon and chlorine The empirical formula is C10Cl12, and the molecular weight545.54 (Hyde 1972; Waters et al 1977; Bell et al 1978; NAS 1978; Menzie 1978; Kaiser 1978)

com-In the United States, the common chemical name is cyclobuta[c,d]pentalene The systematic name is dodecachloropentacyclo 5.3.0.02,6.03,9.04,8decane.Mirex was first prepared in 1946, patented in 1955 by Allied Chemical Company, and introduced

dodecachlorooctahydro-1,3,4-metheno-2H-in 1959 as GC 1283 for use dodecachlorooctahydro-1,3,4-metheno-2H-in pesticidal formulations agadodecachlorooctahydro-1,3,4-metheno-2H-inst hymenopterous dodecachlorooctahydro-1,3,4-metheno-2H-insects, especiallyants It was also marketed under the trade name of Dechlorane for use in flame-retardant coatingsfor various materials Mirex is also known as ENT 25719 (Tucker and Crabtree 1970), CAS 2385-85-5 (Schafer et al 1983), Dechlorane 510, and Dechlorane 4070 (Kaiser 1978) Technical-gradepreparations of mirex consist of 95.19% mirex and less than 2.58 × 10–7% contaminants, mostlykepone C10Cl10O (NAS 1978) Mirex is comparatively soluble in various organic solvents, such asbenzene, carbon tetrachloride, and xylene, with solubilities ranging from about 4000 to 303,000 mg/L.However, mirex has very low solubility in water, not exceeding 1.0 µg/L in freshwater or 0.2 µg/L

in seawater (Bell et al 1978) In biological systems, mirex lipophilicity would account for the highconcentrations observed in fatty tissues and reserves

Mirex, which is composed of 22% carbon and 78% chlorine, is highly resistant to chemical,thermal, and biochemical degradation It is reportedly unaffected by strong acids, bases, and oxidizingagents, and is resistant to photolysis in hydrocarbon solvents, but less so in aliphatic amines Thermaldecomposition begins at about 550˚C and is rapid at 700˚C Degradation products include hexachloro-benzene, hexachlorocyclopentadiene, and kepone Several additional degradation products of mirexhave been isolated, but not all have been identified (Holloman et al 1975; Menzie 1978) At leastone photodegradation product, the 8-monohydro analogue, sometimes accumulates in sediments andanimals, but the fate and effects of these photoproducts are unclear (Cripe and Livingston 1977).Mirex is rapidly adsorbed onto various organic particles in the water column, including algae,and eventually removed to the sediments Not surprisingly, mirex has a long half-life in terrestrialand aquatic sediments; large fractional residues were detected at different locations 12 and 5 yearsafter initial application (Bell et al 1978) Some degradation of mirex to the 10-monohydro analoguewas reported in anaerobic sewage sludge after 2 months in darkness at 30˚C (Menzie 1978) Otherstudies with mirex-contaminated anaerobic soils, anaerobic lake sediments, and soil microorganismsshowed virtually no bacterial degradation over time (Jones and Hodges 1974) In Lake Ontario,mirex from contaminated sediments remained available to lake organisms for many years and, asjudged by present sedimentation rates, mirex may continue to be bioavailable for 200 to 600 years

in that system (Scrudato and DelPrete 1982) Disappearance of mirex from baits over a 12-monthperiod was about 41% for those exposed on the ground, 56% from those exposed in soil, and 84%from those exposed in pond water (de la Cruz and Lue 1978b) Mirex disappearance is probablyrelated to uptake by biological organisms, as has been demonstrated in marine ecosystems con-taminated with mirex (Waters et al 1977), and not to degradation

Mirex is a highly stable chlorinated hydrocarbon with lipophilic properties, and its accumulationand persistence in a wide variety of nontarget biological species has been well documented Thebiological half-life of mirex reportedly ranges from 30 days in quail to 130 days in fish and to morethan 10 months in the fat of female rats (Menzie 1978); this subject area is further developed later

At this juncture, it is sufficient to state that most authorities agree on two points: there is littleevidence of significant mirex metabolism, and mirex ranks among the more biochemically stableorganic pesticides known

21.3.1 Aquatic Organisms

Aquatic organisms are comparatively resistant to mirex in short-term toxicity tests Amongvarious species of freshwater biota, LC50 (96 h) values were not obtained at the highest nominalconcentrations tested of 1000 µg/L for insects, daphnids, and amphipods (Johnson and Finley 1980;Sanders et al 1981) and 100,000 µg/L for five species of fish (Johnson and Finley 1980) Similarresults were reported for other species of freshwater invertebrates (Muncy and Oliver 1963; Lueand de la Cruz 1978) and fishes (Van Valin et al 1968), although waterborne mirex at concentrations

of 1000 µg/L was lethal to postlarval freshwater prawns (Macrobrachium rosenbergerii) in 24 h

(Eversole 1980) It is probable that bioavailable concentrations from the water in each test did notexceed 1.0 µg/L However, delayed mortality frequently occurs for extended periods after exposure,and the potential for adverse effects at the population level remains high (NAS 1978) Latentbiocidal properties of mirex were documented for fish (Van Valin et al 1968; Koenig 1977) andcrustaceans (Ludke et al 1971; Hyde 1972; Cripe and Livingston 1977) Crustaceans were the mostsensitive group examined For example, the crayfish (Procambarus blandingi) immersed in nominalconcentrations of 0.1 to 5.0 µg mirex/L for periods of 6 to 144 h died 5 to 10 days after initialexposure (Ludke et al 1971) Immature crayfish were more sensitive than adults, and mortalitypatterns were similar when mirex was administered in the water or in baits (Ludke et al 1971).

21.3.2 Birds and Mammals

Acute oral toxicity of mirex to warm-blooded organisms was low, except for rats and mice, whichdied 60 to 90 days after treatment with 6 to 10 mg mirex/kg body weight (Table 21.1) Birds werecomparatively resistant The red-winged blackbird (Agelaius phoeniceus) was unaffected at 100 mgmirex/kg body weight, although it was considered the most sensitive of 68 species of birds testedwith 998 chemicals for acute oral toxicity, repellency, and hazard potential (Schafer et al 1983).Mortality due to dietary mirex is variable among species, although high death rates were usuallyassociated with high dietary concentrations and long exposure periods (Table 21.2) One significanteffect of mirex fed to breeding adult chickens, voles, and rats was a decrease in survival of theyoung (Naber and Ware 1965; Shannon 1976; Waters et al 1977; Chu et al 1981) Prairie voles(Micropterus ochrogaster) fed diets containing 15 mg mirex/kg ration bred normally, but all pupsdied by day 21 (Shannon 1976) Survival of the pups of prairie voles decreased in the first litterwhen the diet of the parents contained 10 mg mirex/kg ration, in the second litter when it contained

5 mg/kg, and in the third litter when it contained 0.1, 0.5, 0.7, or 1.0 mg/kg (Shannon 1976)

Table 21.1 Acute Oral Toxicity of Mirex to Birds and Mammals

Dose Organism (mg/kg body weight) Mortality Reference a

Red-winged blackbird, Agelaius phoeniceus

Japanese quail, Coturnix coturnix japonica

a1, Gaines and Kimbrough 1969; 2, Schafer et al 1983; 3, Fujimori et al 1983; 4, Stickel 1963; 5, Hyde 1972;

6, Waters et al 1977; 7, NAS 1978; 8, Schafer et al 1983; 9, Larson et al 1979; 10, Tucker and Crabtree 1970; 11, USPHS 1995.

b Dermal.

• 34 µg/L for fathead minnows, based on impaired reproduction (Buckler et al 1981)

• >34 µg/L for daphnids (Daphnia sp.) and midges (Chaoborus sp.), predicated on daphnid duction and midge emergence (Sanders et al 1981)

repro-Other mirex-induced sublethal effects included reduced photosynthesis in freshwater algae lister et al 1975), gill and kidney histopathology in the goldfish (Carassius auratus) (Van Valin

(Hol-et al 1968), reduced growth in the bluegill (Lepomis macrochirus) (Van Valin et al 1968), cessation

of reproduction in Hydra sp (Lue and de la Cruz 1978), and disrupted behavior in the blue crab(Callinectes sapidus) (Shannon 1976) and the marine annelid (Arenicola cristata) (Schoor andNewman 1976) McCorkle et al (1979) showed that channel catfish (Ictalurus punctatus) areparticularly resistant to high dietary concentrations of mirex; juveniles fed 400 mg mirex/kg rationfor 4 weeks showed no significant changes in enzyme-specific activities of brain, gill, liver, ormuscle However, yearling coho salmon (Oncorhynchus kisutch) fed 50 mg mirex/kg ration for

3 months showed significant reduction in liver weight and whole-body lipid content (Leatherland

Table 21.2 Dietary Toxicity of Mirex to Vertebrate Organisms

Mirex Dietary Concentration Exposure Percent Organism (mg/kg ration) Interval Mortality Reference a

Old-field mouse, Peromyscus polionotus 1.8 60 weeks 20.0 2

Ring-necked pheasant, Phasianus colchicus 1540 5 days 50.0 10

Japanese quail, Coturnix coturnix japonica 5000 5 days 20.0 10

a1, Hyde 1972; 2, Wolfe et al 1979; 3, Chernoff et al 1979; 4, Shannon 1976; 5, Larson et al 1979;

6, Lowe 1982; 7, NAS 1978; 8, Leatherland et al 1979; 9, McCorkle et al 1979; 10, Heath et al 1972.

et al 1979) Additional studies with coho salmon and rainbow trout (Salmo gairdneri) fed 50 mgmirex/kg ration for 10 weeks demonstrated a significant depression in serum calcium, and signif-icant elevation of skeletal magnesium in salmon; trout showed no measurable changes in calciumand magnesium levels in serum, muscle, or skeleton, although growth was reduced, muscle watercontent was elevated, and muscle lipid content was reduced (Leatherland and Sonstegard 1981).Interaction effects of mirex with other anthropogenic contaminants are not well studied, despitethe observations of Koenig (1977) that mixtures of DDT and mirex produced more than additivedeleterious effects on fish survival and reproduction.

21.4.2 Birds

Among captive American kestrels (Falco sparverius) fed 8 mg mirex/kg ration for 69 days byBird et al (1983), there was a marked decline in sperm concentration and a slight compensatoryincrease in semen volume, but an overall net decrease of 70% in sperm number These investigatorsbelieved that migratory raptors feeding on mirex-contaminated food organisms could ingest suffi-cient toxicant to lower semen quality in the breeding season which, coupled with altered courtship,could reduce the fertility of eggs and the reproductive fitness of the individual Altered courtship

in ring-necked doves (Streptopelia capicola) fed dietary organochlorine compounds was reported

by McArthur et al (1983)

Most investigators, however, agree that comparatively high dietary concentrations of mirex hadlittle effect on growth, survival, reproduction, and behavior of nonraptors, including chickens(Gallus sp.), mallards, several species of quail, and red-winged blackbirds For domestic chickens,levels up to 200 mg mirex/kg ration were tolerated without adverse effects on various reproductivevariables (Waters et al 1977), but 300 mg mirex/kg diet for 16 weeks was associated with reducedchick survival, and 600 mg/kg for 16 weeks reduced hatching by 83% and chick survival by 75%(Naber and Ware 1965) Mallard ducklings experienced temporary mild ataxia and regurgitationwhen given a single dose of 2400 mg/kg body weight, but not when given 1200 mg/kg or less(Tucker and Crabtree 1970) Mallards fed diets containing as much as 100 mg mirex/kg ration forprolonged periods showed no significant differences from controls in egg production, shell thick-ness, shell weight, embryonation, hatchability, or duckling survival (Hyde 1972) However, in otherstudies with mallards fed 100 mg mirex/kg diet, eggshells were thinned and duckling survival wasreduced (Waters et al 1977), suggesting that 100 mg mirex/kg ration may not be innocuous tomallards No adverse effects on reproduction were noted in the common bobwhite at 40 mg mirex/kgdiet (Kendall et al 1978), or in two species of quail fed 80 mg mirex/kg ration for 12 weeks (Waters

et al 1977) Red-winged blackbirds were not repelled by foods contaminated with mirex, butconsumed normal rations (Schafer et al 1983); a similar observation was recorded for bobwhites(Baker 1964)

21.4.3 Mammals

Mirex has considerable potential for chronic toxicity because it is only partly metabolized, iseliminated very slowly, and is accumulated in the fat, liver, and brain The most common effectsobserved in small laboratory mammals fed mirex included weight loss, enlarged livers, altered liverenzyme metabolism, and reproductive failure Mirex reportedly crossed placental membranes andaccumulated in fetal tissues Among the progeny of mirex-treated mammals, developmental abnor-malities included cataracts, heart defects, scoliosis, and cleft palates (NAS 1978; Blus 1995).Mirex has caused liver tumors in mice and rats and must be considered a potential humancarcinogen (Waters et al 1977; NAS 1978) Long-term feeding of 50 and 100 mg mirex/kg ration

to rats of both sexes was associated with liver lesions that included neoplastic nodules and tocellular carcinomas; neither sign was found in controls (Ulland et al 1977)

hepa-Adults of selected mammalian species showed a variety of damage effects of mirex:

• Enlarged livers in rats at 25 mg mirex/kg diet (Gaines and Kimbrough 1969) or at a single dose

of 100 mg/kg body weight (Ervin 1982)

• Liver hepatomas in mice at 10 mg mirex/kg body weight daily (Innes et al 1969)

• Decreased incidence of females showing sperm in vaginal smears, decreased litter size, and thyroid histopathology in rats fed 5 mg mirex/kg diet since weaning (Chu et al 1981)

• Elevated blood and serum enzyme levels in rats fed 0.5 mg mirex/kg ration for 28 days (Yarbrough

et al 1981)

• Diarrhea, reduced food and water consumption, body weight loss, decreased blood glucose levels, and disrupted hepatic microsomal mixed function oxidases in mice receiving 10 mg/kg body weight daily (Fujimori et al 1983).

In studies of field mice, decreased litter size was observed at 1.8 mg mirex/kg diet, and completereproductive impairment at 17.6 mg/kg diet after 6 months (Wolfe et al 1979) At comparativelyhigh sublethal mirex concentrations, various deleterious effects were observed: thyroid histopathol-ogy and decreased spermatogenesis in rats fed 75 mg mirex/kg diet for 28 days (Yarbrough et al.1981); abnormal blood chemistry, enlarged livers, reduced spleen size, and loss in body weight ofbeagles fed 100 mg mirex/kg ration for 13 weeks (Larson et al 1979); and decreased hemoglobin,elevated white blood cell counts, reduced growth, liver histopathology, and enlarged livers in ratsfed 320 mg/kg ration for 13 weeks (Larson et al 1979)

Cataract formation, resulting in blindness, in fetuses and pups from maternal rats fed atively low concentrations of dietary mirex is one of the more insidious effects documented Mirexfed to maternal rats at 6 mg/kg body weight daily on days 8 to 15 of gestation, or at 10 mg/kgbody weight daily on days 1 to 4 postpartum, caused cataracts in 50% of fetuses on day 20 ofgestation, and in 58% of pups on day 14 postpartum (Rogers 1982) Plasma glucose levels weredepressed in fetuses with cataracts, and plasma proteins were depressed in neonates; both hypo-proteinemia and hypoglycemia are physiological conditions known to be associated with cataracts(Rogers 1982) Mirex-associated cataractogenicity has been reported in female pups from rats fed

compar-5 mg mirex/kg ration since weaning (Chu et al 1981), in rat pups from females consuming 7 mgmirex/kg ration on days 7 to 16 of gestation or 25 mg/kg diet for 30 days prior to breeding (Chernoff

et al 1979), and in mice fed 12 mg mirex/kg ration (Chernoff et al 1979) Offspring born to treated mothers, but nursed by nontreated mothers showed fewer cataracts (Waters et al 1977).Other fetotoxic effects in rats associated with dietary mirex included:

mirex-• Edema and undescended testes (Chernoff et al 1979)

• Lowered blood plasma proteins, and heart disorders, including tachycardia and blockages (Grabowski 1981)

• Hydrocephaly; decreases in weight of brain, lung, liver, and kidney; decreases in liver glycogen, kidney proteins and alkaline phosphatase; and disrupted brain DNA and protein metabolism (Kav- lock et al 1982)

In prairie voles exposed continuously to dietary mirex of 0.5, 0.7, 1.0, 5.0, or 10.0 mg/kg ration,the numbers of litters produced decreased (Shannon 1976) Maximum number of litters per yearwere four at 1.0 mg mirex/kg ration, three at 5.0 mg/kg, and two at 10.0 mg/kg ration Furthermore,the number of offspring per litter also decreased progressively Concentrations as low as 0.1 mgmirex/kg ration of adults were associated with delayed maturation of pups and with an increase innumber of days required to attain various behavioral plateaus such as bar-holding ability, hind-limbplacing, and negative geotaxis (Shannon 1976) On the basis of residue data from field studies, as

is shown later, these results strongly suggest that mirex was harmful to the reproductive performanceand behavioral development of prairie voles at environmental levels approaching 4.2 g mirex/ha,

a level used to control fire ants before mirex was banned

21.5 BIOACCUMULATION

21.5.1 Aquatic Organisms

All aquatic species tested accumulated mirex from the medium and concentrated it over ambientwater levels by factors ranging up to several orders of magnitude Uptake was positively correlatedwith nominal dose in the water column (Table 21.3) Other investigators have reported bioconcen-tration factors from water of 8025 in daphnids (Sanders et al 1981), 12,200 in bluegills (Skaar

et al 1981), 56,000 in fathead minnows (Huckins et al 1982), and 126,600 in the digestive gland

of crayfish (Ludke et al 1971) Rapid uptake of mirex by marine crabs, shrimps, oysters, killifishes,and algae was reported after the application of mirex baits to coastal marshes (Waters et al 1977;Cripe and Livingston 1977) Mirex was also accumulated from the diet (Table 21.3) (Ludke et al.1971; Zitko 1980), but not as readily as from the medium Dietary bioaccumulation studies withguppies and goldfish show that mirex and other persistent hydrophobic chemicals are retained inthe organism and biomagnify through food chains because of their hydrophobicity (Gobas et al

1989, 1993; Clark and Mackay 1991) Mirex may also be accumulated from contaminated sediments

by marine teleosts (Kobylinski and Livingston 1975), but such accumulation has not been lished conclusively Although terrestrial plants, such as peas and beans, accumulate mirex at fieldapplication levels, mangrove seedlings require environmentally high levels of 11.2 kg mirex/habefore accumulation occurs (as quoted in Shannon 1976)

estab-There is general agreement that aquatic biota subjected to mirex-contaminated environmentscontinue to accumulate mirex, and that equilibrium is rarely attained before death of the organismfrom mirex poisoning or from other causes There is also general agreement that mirex resistsmetabolic and microbial degradation, exhibits considerable movement through food chains, and ispotentially dangerous to consumers at the higher trophic levels (Hollister et al 1975; NAS 1978;Mehrle et al 1981; Eisler 1985) Marine algae, for example, showed a significant linear correlationbetween amounts accumulated and mirex concentrations in the medium If a similar situation existed

in nature, marine unicellular algae would accumulate mirex and, when grazed upon, act as passivetransporters to higher trophic food chain compartments (Hollister et al 1975) The evidence forelimination rates of mirex from aquatic biota on transfer to mirex-free media is not as clear.Biological half-times of mirex have been reported as 12 h for daphnids (Sanders et al 1981), morethan 28 days for fathead minnows (Huckins et al 1982), about 70 days in Atlantic salmon (Salmo salar) (Zitko 1980), 130 days for mosquitofish (Gambusia affinis), and 250 days for pinfish (asquoted in Skea et al 1981) However, Skea et al (1981) averred that biological half-times may bemuch longer if organism growth is incorporated into rate elimination models For example, brooktrout (Salvelinus fontinalis) fed 29 mg mirex/kg ration for 104 days contained 6.3 mg/kg bodyweight or a total of 1.1 mg of mirex in whole fish At day 385 postexposure, after the trout hadtripled in body weight, these values were 2.1 mg/kg body weight, an apparent loss of 67%; however,

on a whole-fish basis, trout contained 1.2 mg, thus showing essentially no elimination on a organism basis (Skea et al 1981)

total-No mirex degradation products were detected in whole fathead minnow or in hydrosoils underaerobic or anaerobic conditions (Huckins et al 1982) In contrast, three metabolites were detected

in coastal marshes after mirex bait application, one of which, photomirex, was accumulated by fishand oysters (Cripe and Livingston 1977) The fate and effects of mirex photoproducts in theenvironment are unclear and merit additional research

The significance of mirex residues in various tissues is unresolved, as is the exact mode ofaction of mirex and its metabolites Minchew et al (1980) and others indicated that mirex is aneurotoxic agent, with a mode of action similar to that of other chlorinated hydrocarbon insecticides,such as DDT In studies with crayfish and radiolabeled mirex, mirex toxicosis was associated withneurotoxic effects that included hyperactivity, uncoordinated movements, loss of equilibrium, and

paralysis (Minchew et al 1980) Before death, the most significant differences in mirex distributions

in crayfish were the increases in concentrations in neural tissues, such as brain and nerve cord, byfactors up to 14 (or 0.4 mg/kg) in low-dose groups held in solutions containing 7.4 µg mirex/L,and up to 300 (or 6.2 mg/kg) in high-dose groups held in solutions with 74.0 µg/L With continuedexposure, levels in the green gland and neural tissues approached the levels in the hepatopancreasand intestine (Table 21.3) Schoor (1979) also demonstrated that mirex accumulates in the crusta-cean hepatopancreas, but suggested that other tissues, such as muscle and exoskeleton, have specificbinding sites that, once filled, shunt excess mirex to hepatopancreas storage sites

21.5.2 Birds and Mammals

Like aquatic organisms, birds and mammals accumulated mirex in tissue lipids, and the greateraccumulations were associated with the longer exposure intervals and higher dosages (Table 21.3).Sexual condition of the organism may modify bioconcentration potential For example, in adiposefat of the bobwhite, males contained 10 times dietary levels and females only 5 times dietary levels;the difference was attributed to mirex loss through egg laying (Kendall et al 1978)

Data on excretion kinetics of mirex are incomplete Prairie voles fed mirex for 90 days containeddetectable whole-body levels 4 months after being placed on a mirex-free diet (Shannon 1976).Levels of mirex in voles after 4 months on uncontaminated feed were still far above levels in theirmirex diets Humans living in areas where mirex has been used for ant control contained 0.16 to5.94 mg/kg in adipose fat; 60% of the mirex was excreted and most of the rest was stored in bodytissues, especially fat (28%), and in lesser amounts of 0.2 to 3% in muscle, liver, kidney, andintestines (Waters et al 1977) Almost all excretion of mirex takes place through feces; less than1% is excreted in urine and milk The loss rate pattern is biphasic, the fast phase was estimated at

38 h and the slow phase at up to 100 days Mirex binds firmly to soluble liver proteins and appears

to be retained in fatty tissues, a property that may contribute to its long biological half-life Chickensgiven single doses of mirex at 30 mg/kg intravenously or 300 mg/kg orally demonstrated a biphasicdecline in blood concentrations (Ahrens et al 1980) The fast component, constituting about 25%

of the total, was lost during the first 24 h; the loss of the slow component was estimated to be at

a constant rate of about 0.03% daily, suggesting a half-life of about 3 years Growing chicks fed

1 or 10 mg/kg dietary mirex for 1 week lost the compound rather rapidly; disappearance half-timeswere 25 days for skin and 32 days for fat (Ahrens et al 1980) It is clear that much additionalresearch is warranted on loss rate kinetics of this persistent compound and its metabolites

Table 21.3 Uptake of Mirex from Ambient Medium or Diet by Selected Species

Habitat, Organism,

and Tissue

Mirex in Medium (M) ( g/L) or in Diet (D) (mg/kg) Exposure

Bioconcentration factor (BCF) Reference a AQUATIC, FRESHWATER

Bluegill, Lepomis macrochirus

of mirex toxicity)

Bioconcentration factor (BCF) Reference a

BIRDS AND MAMMALS

Mallards, Anas platyrhynchos

(exposed adults)

American kestrels, Falco

sparverius, yearling males

Common bobwhite, Colinus

virginianus

Mammals

Rat, Rattus sp.

a1, Buckler et al 1981; 2, Van Valin et al 1968; 3, Zitko 1980; 4, Skea et al 1981; 5, Minchew et al 1980;

6, Koenig 1977; 7, Kobylinski and Livingston 1975; 8, Lee et al 1975; 9, Schoor 1979; 10, Hollister et al 1975;

11, Waters et al 1977; 12, Ahrens et al 1980; 13, Hyde 1972; 14, Bird et al 1983; 15, Kendall et al 1978;

16, NAS 1978; 17, Chu et al 1981; 18, Wolfe et al 1979.

Table 21.3 (continued) Uptake of Mirex from Ambient Medium or Diet by Selected Species

Habitat, Organism,

and Tissue

Mirex in Medium (M) ( g/L) or in Diet (D) (mg/kg) Exposure

Bioconcentration factor (BCF) Reference a

21.6 MIREX IN THE SOUTHEASTERN UNITED STATES

Between 1961 and 1975, about 400,000 kg mirex were used in pesticidal formulations, of which

approximately 250,000 kg were sold in the southeastern United States for control of native and

imported fire ants (Solenopsis spp.) Most of the rest was exported to Brazil for use in fire ant

control in that country (NAS 1978) Mirex was also used to control big-headed ant populations in

Hawaiian pineapple fields (Bell et al 1978), Australian termites (Paton and Miller 1980), South

American leaf cutter ants, South African harvester termites, and, in the United States, western

harvester ants and yellow jackets (Shannon 1976) Chemical control measures for imported fire

ants began in the southeastern United States during the 1950s with the use of heptachlor, chlordane,

and dieldrin The large mounds built by ants in cultivated fields were believed to interfere with

mowing and harvesting operations; the “vicious sting” of the insects presented a hazard to workers

harvesting the crops; and the species was considered to be a pest in school playgrounds and homes

(Lowe 1982) In 1965, the use of organochlorine insecticides to control fire ants was discontinued,

due partly to their high acute toxicity to nontarget biota and their persistence Previously used

compounds were replaced by mirex 4X bait formulations, consisting of 0.3% mirex by weight,

dissolved in 14.7% soybean oil, and soaked into corncob grits (85%) Initially, the 4X baits were

broadcast from low-flying airplanes at a total yearly rate of 1.4 kg bait/ha (1.25 lb total bait/acre)

or 4.2 g mirex/ha Usually, three applications were made yearly More than 50 million ha in nine

southeastern states were treated over a 10-year period Later, dosages were modified downward,

and mirex was applied to mounds directly Ecologically sensitive areas, such as estuaries, prime

wildlife habitats, heavily forested areas, and state and federal parks, were avoided In 1977, for

example, the formulation was changed to 0.1% mirex and the application rate lowered to 1.12 g/ha;

about 8200 kg of the lower-concentration bait were manufactured in 1977 (Bell et al 1978) Under

ideal aerial application conditions, about 140 particles of mirex-impregnated bait were distributed

per square meter When an infested area is treated, the bait is rapidly scavenged by the oil-loving

fire ant workers, placed in the mound, and distributed throughout the colony, including queen and

brood, before any toxic effects become evident Death occurs in several days to weeks The exact

mode of action is unknown, but is believed to be similar to that of other neurotoxic agents such as

DDT (Waters et al 1977; NAS 1978)

Widespread use of mirex may lead to altered population structure in terrestrial systems, with

resurgence or escalation of nontarget pests due to selective mirex-induced mortality of predators

(NAS 1978) For example, populations of immature horn flies and rove beetles, two species of

arthropods normally preyed upon by fire ants, were higher in mirex-treated areas than in control

areas (Howard and Oliver 1978) Conversely, other species, such as crickets, ground beetles, and

various species of oil-loving ants, were directly affected and populations were still depressed or

eliminated 14 months posttreatment (NAS 1978), whereas fire ants recovered to higher than

pretreatment levels, as judged by mound numbers and mound size (Summerlin et al 1977)

Field results from aquatic and terrestrial ecosystems receiving mirex bait formulations indicated,

with minor exceptions, that mirex accumulates sequentially in food complexes and concentrates in

animals at the higher trophic levels In both ecosystems, omnivores and top carnivores contained

the highest residues (Hyde 1972; Shannon 1976; Waters et al 1977; de la Cruz and Lue 1978a;

Hunter et al 1980; Eisler 1985) In South Carolina, where the 4X formulation was used to control

fire ants from 1969 to 1971, mirex was translocated from treated lands to nearby marshes and

estuarine biota, including crustaceans, marsh birds, and raccoons (Lowe 1982) Juvenile marine

crustaceans showed delayed toxic effects after ingesting mirex baits, or after being exposed to low

concentrations in seawater About 18 months posttreatment, mirex residues of 1.3 to 17.0 mg/kg

were detected in shrimp, mammals, and birds (Table 21.4); however, 24 months after the last mirex

treatment, less than 10% of all samples collected contained detectable residues (Lowe 1982) A

similar study was conducted in pasturelands of bahia grass (Paspalium notatum) (Markin 1981)

Within a month after application, the target fire ant colonies were dead Of the 4.2 g mirex/ha

applied to the 164 ha block, 100% was accounted for on day 1, 63% at 1 month, and 3% at 1 year

(Table 21.5) Unaccounted mirex residues could include loss through biodegradation; through

movement out of the study area by migratory insects, birds, other fauna, and groundwater; and

through photodecomposition and volatilization (Markin 1981)

Mirex residues in bobwhites from a South Carolina game management area were documented

after treatment with 4.2 g mirex/ha (Kendall et al 1977) Pretreatment residues in bobwhites ranged

from nondetectable to 0.17 mg mirex/kg breast muscle on a dry-weight basis, and 0.25 to 2.8 mg/kg

in adipose tissues on a lipid-weight basis Mirex residues in adipose tissue increased up to 5 times

within 1 month posttreatment and declined thereafter; however, another residue peak was noted in

the spring after mirex treatment and corresponded with insect emergence (Kendall et al 1977)

Mirex concentrations in muscle and liver of mammalian wildlife in Alabama and Georgia during

the period 1973 to 1976 from reference areas were always less than 0.04 mg mirex/kg FW in

muscle and less than 0.07 mg/kg FW in liver (Hill and Dent 1985) In mirex-treated areas,

conspecifics were collected up to 2 years posttreatment Maximum concentrations of mirex in

muscle and liver from mirex-treated areas were always less than 1.0 mg/kg FW in raccoons, bobcats

(Lynx rufus), mink (Mustela vison ), and foxes (Urocyon sp., Vulpes sp.) Higher concentrations of

3.7 mg/kg FW in muscle and 1.1 mg/kg FW in liver were measured in the river otter (Lutra

Table 21.4 Mirex Residues in Water, Sediments, and Fauna

in a South Carolina Coastal Marsh 18 months after Application of 4.2 g/ha

Modified from Lowe, J.I 1982 Mirex, fire ants, and estuaries Pages

63-70 in Proceedings of the Workshop on Agrichemicals and rine Productivity Duke Univ Mar Lab., Beaufort, NC Sept 18-19,

Estua-1980 U.S Dep Comm NOAA/OMPA.

Table 21.5 Temporal Persistence of Residues for 1 Year after Applications

of Mirex 4X Formulation to Bahia Grass Pastures (Values represent rounded percentages recovered of the original 4.2 g/ha applied.)

Time, postapplication Sample 1 d 2 wk 1 mo 3 mo 6 mo 9 mo 12 mo

a Grit now included with pasture soil.

b Mounds badly weathered, not possible to identify.

Modified from Markin, G.P 1981 Translocation and fate of the insecticide mirex

within a bahia grass pasture ecosystem Environ Pollut 26A:227-241.

canadensis) 1 year posttreatment, 3.5 mg/kg FW in muscle of skunks (Spilogale sp., Mephitis sp.)

6 months posttreatment, and 1.1 to 1.5 mg/kg FW in embryos and muscle of the opossum phius marsupialis) 6 to 12 months after treatment (Hill and Dent 1985).

(Didel-Heavily treated watershed areas in Mississippi were investigated by Wolfe and Norment (1973)and Holcombe and Parker (1979) After treatment, mirex residues were elevated in crayfish andstream fish Among mammals, residues were highest in carnivores and insectivores, lower inomnivores, and lowest in herbivores (Wolfe and Norment 1973) Mirex residues in liver and eggs

were substantially higher in the box turtle (Terrapene carolina), an omnivorous feeder, than in the herbivorous slider turtle (Chrysemys scripta); mirex did not accumulate for protracted periods in

tissues of these comparatively long-lived reptiles (Holcombe and Parker 1979) Among migratory

reptiles, mirex was detected in only 11% of the eggs of the loggerhead turtle (Caretta caretta) and not at all in eggs of the green turtle (Chelonia mydas) collected during summer 1976 in Florida

(Clark and Krynitsky 1980) However, DDT or its isomers were present in all eggs of both species,and PCBs were detected in all loggerhead turtle eggs The low levels of mirex and other orga-nochlorine contaminants suggest that these turtles, when not nesting, live and feed in areas remotefrom Florida lands treated with mirex and other insecticides (Clark and Krynitsky 1980)

A 10-5 bait formulation containing 0.1% mirex was designed to make more of the toxicantavailable to the fire ant and less to nontarget biota In one study, the 10-5 formulation was applied to

a previously untreated 8000-ha area near Jacksonville, Florida, infested with fire ants (Wheeler et al.1977) The bait was applied by airplane at 1.12 kg/ha, or 1.12 g mirex/ha Insects accumulated mirex

to the greatest extent during the first 6 months after application, and most of the mirex was lost by

12 months (Table 21.6) Other invertebrates accumulated only low levels during the first 9 months,and no residues were detected after 12 months Fish also showed low concentrations for 9 monthsand no detectable residues afterward Amphibians contained detectable residues after 12 months, butnot at 24; and reptiles contained measurable, but low, residues for the entire 24-month study period.Mammals had higher residue levels than reptiles, particularly in fat, whereas birds contained low tomoderate residues (Table 21.6) After 24 months, mirex was found infrequently and only at lowconcentrations in birds, mammals, reptiles, and insects It was concluded that 10-5 mirex formulationswere as effective in controlling fire ants as the 4X formulation and that residues in nontarget specieswere reduced from that following 4X treatment, or were lacking (Wheeler et al 1977)

Eggs of the American crocodile (Crocodylus acutus) from the Florida Everglades contained up

to 2.9 mg/kg fresh weight of DDE and 0.86 mg/kg of polychlorinated biphenyls, but less than

0.02 mg mirex/kg (Hall et al 1979) Livers of the deep sea fish (Antimora rostrata) collected from

1971 to 1974 from a depth of 2500 m off the U.S east coast, contained measurable concentrations

of DDT and its degradation products, and dieldrin, but no mirex (Barber and Warlen 1979)

Table 21.6 Mirex Residues in Fauna near Jacksonville, Florida,

at Various Intervals Posttreatment Following Single Application of 1.12 g mirex/ha

Taxonomic Group and Time Maximum Residue (months posttreatment) (mg/kg wet weight whole organism) INSECTS

21.7 MIREX IN THE GREAT LAKES

Between 1959 and 1975, 1.5 million kg mirex were sold, of which 74%, or more than 1.1 million

kg, were predominantly Dechlorane, a compound used in flame-resistant polymer formulations ofelectronic components and fabrics (Bell et al., 1978; NAS 1978) The total amounts are only approx-imate because almost half the mirex sold from 1962 to 1973 could not be accounted for (NAS 1978).Mirex loadings to Lake Ontario were estimated at 200 kg per year in 1960 to 1962, which decreased

to 28 kg in 1980 (Halfon 1987) Mirex entered Lake Ontario mainly from the Niagara and OswegoRivers About 700 kg mirex were present in the bottom sediments of Lake Ontario in 1968, 1600 kg

in 1976, and 1784 kg in 1981 (Halfon 1984) Kaiser (1978) reported that all fish species in LakeOntario were contaminated with mirex, and that concentrations in half the species exceeded the Foodand Drug Administration guideline of 0.1 mg/kg; other aquatic species had mirex residues near this

level Reproduction of the herring gull (Larus argentatus) on Lake Ontario was poor; mirex levels

were an order of magnitude higher in gull eggs from Lake Ontario than in eggs from other GreatLakes locations (Kaiser 1978) It was concluded that the probable source of contamination was achemical manufacturer that used mirex (Dechlorane) as a flame retardant, and that only Lake Ontariowas contaminated (Kaiser 1978; NAS 1978) Until 1988, mirex had been reported for only a fewlocations in the Great Lakes, primarily Lake Ontario and the St Lawrence River Since 1988, however,mirex in water and fish samples has been measured from the other Great Lakes (Sergeant et al 1993).Gilman et al (1977, 1978) observed poor reproductive success and declines in colony size ofthe herring gull at Lake Ontario at a time when dramatic increases of this species were reportedalong the Atlantic seaboard In 1975, herring gull reproduction in Lake Ontario colonies was aboutone tenth that of colonies on the other four Great Lakes In addition, in Lake Ontario colonies,there were reductions in nest site defense, the number of eggs per clutch, hatchability of eggs, andchick survival Hatching success of Lake Ontario gull eggs was 23 to 26%, compared with 53 to79% for eggs from other areas Analysis of herring gull eggs from all colonies for organochlorinecompounds and mercury demonstrated that eggs from Lake Ontario colonies had mean mirex levels

of 5.06 mg/kg fresh weight (range, 2.0 to 18.6), or about 10 times more mirex than any othercolony Mean PCB and mercury levels were up to 2.8 and 2.3 times higher, respectively, in gulleggs from Lake Ontario than in those from other colonies, but only mirex levels could account for

Modified from Wheeler, W.B., D.P Jouvenay, D.P Wojcik, W.A Banks, C.H.

Van Middelem, C.S Lofgren, S Nesbitt, L Williams, and R Brown 1977.

Mirex residues in nontarget organisms after application of 10-5 bait for fire

ant control, Northeast Florida — 1972–74 Pestic Monitor Jour 11:146-156.

Table 21.6 (continued) Mirex Residues in Fauna near Jacksonville,

Florida, at Various Intervals Posttreatment Following Single Application of 1.12 g mirex/ha

Taxonomic Group and Time Maximum Residue (months posttreatment) (mg/kg wet weight whole organism)

the colony declines (Gilman et al 1977, 1978) Short-term deviations from long-term trends inmirex concentrations in eggs of herring gulls from Lake Ontario seem to be correlated with weatherpatterns (i.e., warm spring weather conducive to phytoplankton growth produces relatively uncon-taminated plankton, which results in less contamination for gulls during the critical period of eggyolk formation — and the reverse for cold spring weather) (Smith 1995) As judged by log-linearregression models, the half-life for mirex in herring gull eggs was 1.9 to 2.1 years, or essentiallynone was lost during egg incubation (Weseloh et al 1979) Reproductive success of the LakeOntario herring gull colonies improved after the early 1970s, an improvement that was directlyparalleled by a decline in mirex, other organochlorine pesticides, and PCBs (Weseloh et al 1979).

Concentrations of mirex and other contaminants in eggs of the Caspian tern (Sterna caspia)

from the Great Lakes are declining, and tern populations are increasing (Struger and Weseloh 1985;Ewins et al 1994) In Lake Huron, mirex concentrations in Caspian tern eggs declined from0.51 mg/kg FW in 1976 to 0.12 mg/kg FW in 1991, equivalent to a decline of 8.6% annually InLake Ontario, mirex concentrations in tern eggs declined from 1.6 mg/kg FW in 1981 to 0.77 mg/kg

FW in 1991, a decline of 7.1% annually (Ewins et al 1994) Similar trends are reported for eggs

of herring gulls from Lakes Michigan, Huron, and Ontario (Ewins et al 1992, 1994), whole lake

trout (Salvelinus namaycush) from Lake Ontario (Borgmann and Whittle 1991), and whole of-the-year spottail shiners (Notropis hudsonicus) throughout the Great lakes (Suns et al 1993).

young-The fate of mirex in the environment and the associated transfer mechanisms have not beenwell defined (NAS 1978) One of the more significant works on this subject area was that byNorstrom et al (1978), who documented levels of mirex and its degradation products in herringgull eggs collected from Lake Ontario in 1977 (Table 21.7) They concluded that photodegradationwas the only feasible mechanism for production of the degradation compounds, although mirexand its photoproducts rapidly become sequestered in the ecosystem and protected from furtherdegradation Norstrom et al (1980) found mirex degradation products in herring gull eggs from all

of the Great Lakes and suggested that a high proportion of mirex and related compounds in herringgull eggs from Lakes Erie and Huron originated from Lake Ontario fish, whereas lower levels ineggs from Lakes Superior and Michigan originated from other sources Mirex in sediments wasconsidered an unlikely source because it was not being recycled into the ecosystem at an appreciable

rate (Norstrom et al 1980) Migrating salmon (Oncorhynchus spp.) make a significant contribution

to the upstream transport of mirex from Lake Ontario, estimated at 53 to 121 g mirex annually(Lewis and Makarewicz 1988; Scrudato and McDowell 1989) Ingestion of salmon eggs by browntrout, decomposition of salmon carcasses by blowfly larvae, and ingestion of carcasses by aquaticand terrestrial scavengers are all means by which mirex is introduced to upstream environments(Scrudato and McDowell 1989) A harvest rate of 50% by fisherman represents a removal of anadditional 61 g mirex annually from Lake Ontario (Lewis and Makarewicz 1988)

Table 21.7 Mirex and Its Degradation Products in Herring Gull Eggs Collected

from the Great Lakes in 1977

Mirex Concentration, Percent of Samples Compound (mg/kg fresh weight) Containing Compound

Modified from Norstrom, R.J., D.J Hallett, F.I Onuska, and M.E Comba 1980 Mirex and its

degradation products in Great Lakes herring gulls Environ Sci Technol 14:860-866.

Biomagnification of mirex through food chains was investigated by Norstrom et al (1978).

Their basic assumption was that both herring gulls and coho salmon ate alewives (Alosa

pseudoharengus) and rainbow smelt (Osmerus mordax) Mirex residues in these organisms, in

mg/kg (parts per million) fresh weight, were 4.4 in gull eggs, 0.23 in salmon muscle, 0.10 in salmon

liver, and 0.09 in whole alewives and smelt retrieved from stomachs of salmon Bioconcentration

factors (BCFs) from prey to predator ranged up to 50, and those from water to gull egg were

estimated to be near 25 million (Table 21.8) Norstrom et al (1978) indicated that salmon muscle

and gull eggs are complementary indicators of organochlorine contamination in the Great Lakes

Among Great Lakes fishes, the highest mirex value recorded was 1.39 mg/kg FW in whole

American eels (Anguilla rostrata) collected from Lake Ontario and was substantially in excess of

the tolerated limit of 0.3 mg/kg FW for human consumption at that time (NAS 1978) In the early

1980s, mirex was detected in 100% of the American eels sampled from Lake Ontario (Dutil et al

1985) High mirex values were also reported in chinook salmon (Oncorhynchus tshawytscha) and

coho salmon (Oncorhynchus kisutch) from South Sandy Creek, a tributary of Lake Ontario, during

autumn 1976 As a consequence, possession of all fish from that area was prohibited by the State

of New York (Farr and Blake 1979) Mirex concentrations in coho and chinook salmon tissues from

Lake Ontario in 1977/78 ranged between 0.07 and 0.24 mg/kg FW tissue and increased with

individual fish weight in direct relation to lipid content (Insalaco et al 1982) The significance of

mirex residues in salmonid fishes is unclear Skea et al (1981), in laboratory studies with brook

trout, showed that whole-body residues of 6.3 mg/kg fish weight were not associated with adverse

effects on growth or survival and speculated that long-lived species, such as the lake trout, would

probably continue to accumulate mirex in Lake Ontario as long as they were exposed, and may

continue to contain residues for most of their lives, even after the source has been eliminated

There was no widespread mirex contamination of urban environments near Lake Ontario as a

result of Dechlorane use, although local contamination of the Lake Ontario area was high when

compared with other Great Lakes areas (NAS 1978) Among humans living in the Great Lakes

area, there was great concern that mother’s milk might be contaminated, owing to the high

lipophilicity of mirex Bush (1983) found mirex concentrations in mother’s milk from residents of

New York state to be 0.07 µg/L in Albany, 0.12 µg/L in Oswego, and 0.16 µg/L in Rochester,

confirming that mirex was present in human milk but that concentrations were sufficiently low to

be of little toxicological significance It is noteworthy that none of the mothers had eaten Lake

Ontario fish or any freshwater fish, and only a few had eaten marine fishes (Bush 1983) For a

5-kg infant consuming 500 g milk daily, this amount would approximate a daily dietary intake of

0.01 µg mirex/kg body weight (Bush 1983), or about 1/10,000 of the lowest recorded dietary value

causing delayed maturation in prairie voles (Shannon 1976) It is not known if a safety factor of

10,000 is sufficient to protect human health against delayed toxic effects of mirex, but it now

appears reasonable to believe that it is

Table 21.8 Biomagnification of Mirex in Great Lakes Food Chains

Bioconcentration

Water Whole rainbow smelt (Osmerus mordax) or

whole alewife (Alosa pseudoharengus)

500,000 Water Muscle of coho salmon (Oncorhynchus kisutch) 1,500,000

Modified from Norstrom, R.J., D.J Hallett, and R.A Sonstegard 1978 Coho salmon (Oncorhynchus

kisutch) and herring gulls (Larus argentatus) as indicators of organochlorine contamination in Lake

Ontario Jour Fish Res Board Canada 35:1401-1409.

21.8 MIREX IN OTHER GEOGRAPHIC AREAS

Mirex residues were determined in birds collected nationwide or from large geographic areas

of the United States; however, aside from the Southeast and the Great Lakes, concentrations werelow, considered nonhazardous, and occurred in a relatively small proportion of the samples collected(Cain and Bunck 1983; Wood et al 1996) Among wings of mallards and American black ducks

(Anas rubripes) collected from the four major flyways during 1976/77, mirex concentrations were

highest and percent occurrence greatest in samples from the Atlantic Flyway: mallards, 50%occurrence, 0.14 mg/kg fresh weight; black ducks, 19% and 0.04 mg/kg (White 1979) Data formallards collected from other flyways follow: Mississippi, 29% and 0.03 mg/kg; Central, 14% and0.06 mg/kg; and Pacific 4% and 0.03 mg/kg (White 1979) Carcasses of several species of heronsfound dead or moribund nationwide from 1966 to 1980 were analyzed for a variety of commonorganochlorine pesticides by Ohlendorf et al (1981) They detected mirex in less than 15% of thecarcasses, a comparatively low frequency, and only in nonhazardous concentrations However, about20% of all herons found dead or moribund had lethal or hazardous concentrations of dieldrin or

DDT In bald eagles (Haliaeetus leucocephalus) found dead nationwide, elevated mirex levels were

recorded in carcass lipids (24.0 mg/kg) and in fresh brain tissues (0.22 mg/kg) (Barbehenn andReichel 1981) Among endangered species such as the bald eagle, it was determined that the mostreliable indicator for assessing risk of organochlorine compounds was the ratio of carcass to brainresidues on a lipid weight basis (Barbehenn and Reichel 1981) Wings from American woodcocks

(Philohela minor) collected from 11 states in 1970/71 and 14 states in 1971/72 were analyzed for

mirex and other compounds by McLane et al (1978) Mirex residues in the 1971/72 wings showedthe same geographical pattern of recovery as those observed in 1970/71: residues were highest inthe southern states and New Jersey, and lowest in the northern and midwestern states Mirex residueswere significantly lower in 1971/72 than in 1970/71 As judged by the analysis of wings of immaturewoodcocks in Louisiana, mirex residues were significantly lower in immatures than in adults:2.48 mg/kg lipid weight vs 6.20 mg/kg, respectively (McLane et al 1978)

Mirex concentrations in bald eagle eggs collected nationwide between 1969 and 1979 rangedfrom 0.03 to 2.0 mg/kg FW, and were highest in Florida and the Chesapeake Bay region (Wiemeyer

et al 1984) Up to 87% of bald eagle eggs from Florida and the Chesapeake Bay had detectablemirex residues, whereas this value was as low as 17% in Alaska Wiemeyer et al (1984) note thateggs from successful bald eagle nests had 0.03 mg mirex/kg FW and lower, but eggs from

unsuccessful nests had 0.05 mg/kg FW and higher Eggs of Cooper’s hawk, Accipiter cooperi,

collected in 1980 from various locations, all contained more than 0.05 mg mirex/kg FW trations were highest in Pennsylvania (with 0.84 mg/kg FW) and Wisconsin (with 1.6 mg/kg FW)

Concen-(Pattee et al 1985) Eggs of the loggerhead shrike (Lanius ludovicianus) from the Shenandoah

Valley region of Virginia in 1985/86 contained an average of 0.04 mg mirex/kg FW, with a 63%frequency of occurrence; loggerhead shrike populations in that region are declining but the cause

of the decline is not known with certainty (Blumton et al 1990) Eggs of the ring-necked grebe

(Podiceps grisigena) from Manitoba, Canada, in 1980/81, had as much as 28.6 mg mirex/kg lipid

weight, and this may account, in part, for the high nesting loss of 79% observed in grebes at that

time (De Smet 1987) Mirex and other organochlorine compounds in eggs of anhingas (Anhinga anhinga) and 17 species of waders (including herons, egrets, bitterns, ibises, and storks) were

measured in various locations throughout the eastern United States during 1972 and 1973 dorf et al 1979) The highest mean concentration of 0.74 mg mirex/kg, range 0.19 to 2.5 mg/kg,

(Ohlen-was found in eggs of the green heron (Butorides striatus) from the Savannah National Wildlife Refuge in South Carolina; a single egg of the cattle egret (Bubulucus ibis) analyzed from there

contained 2.9 mg mirex/kg However, the overall frequency of mirex occurrence was higher in eggscollected from the Great Lakes region (24%) than in those from the South Atlantic coast (15.6%),

inland areas (10.7%), Gulf Coast (4.4%), or North Atlantic region (3.2%) Measurable mirexresidues were detected in migratory birds collected from a variety of locations, including areas farfrom known sources or applications of mirex For example, 22% of all eggs from 19 species ofAlaskan seabirds collected in 1973 to 1976 contained mirex The highest concentration was

0.044 mg/kg in eggs of a fork-tailed storm petrel (Oceanodroma furcata) from the Barren Islands.

Mirex residues were low compared with those of other organochlorine compounds (Ohlendorf et al

1982) Eggs from the clapper rail (Rallus longirostris) collected in New Jersey from 1972 to 1974

contained 0.16 to 0.45 mg mirex/kg (Klaas et al 1980) Eggs from the greater black-backed gull

(Larus marinus) collected from Appledore Island, Maine, in 1977 contained up to 0.26 mg/kg, but

no mirex was detected in eggs of common eider (Somateria mollissima) or herring gull from the

same area (Szaro et al 1977) The greater black-backed gull is an active carnivore; 36 to 52% ofits diet consists of small birds and mammals, whereas these items compose less than 1% in eiderand herring gull diets The higher mirex levels in black-backed gulls are attributed to its predatory

feeding habits (Szaro et al 1979) In New England, eggs of the black-crowned night-heron ticorax nycticorax) contained between 0.28 and 0.66 mg mirex/kg wet weight in 1973; in 1979,

(Nyc-this range was 0.11 to 0.37 mg/kg (Custer et al 1983) Falcon eggs contained detectable mirex;

levels were highest in the pigeon hawk (Falco columbarius) (0.25 mg/kg) and in the peregrine falcon (Falco peregrinus) (0.43 mg/kg), two species that feed on migratory birds or migrate to mirex-impacted areas (Kaiser 1978) Active mirex was also found in eggs of a cormorant (Phala- crocorax sp.) from the Bay of Fundy on the Atlantic coast; the suspected source of contamination

was the southern wintering range (Kaiser 1978)

Mirex residues in 20 great horned owls (Bubo virginianus) found dead or dying in New York

state in 1980 to 1982 contained concentrations of mirex and PCBs higher than those reported forgreat horned owls elsewhere (Stone and Okoniewski 1983) Owls in “poor flesh” contained higherresidues than those in “good flesh”; these values were 6.3 mg/kg FW vs 0.07 mg/kg FW for brain,and 5.6 mg/kg FW vs 0.1 mg/kg FW for liver (Stone and Okoniewski 1983) Waterfowl collectedfrom upstate New York between 1979 and 1982 had about 0.07 mg mirex/kg FW breast muscleand 0.28 mg/kg FW subcutaneous fat (Kim et al 1984, 1985)

Mink (Mustela vison) collected from the Northwest Territories of Canada between 1991 and

1995 had liver mirex concentrations between 0.08 and 0.39 µg/kg FW These extremely low mirexconcentrations were, nevertheless, higher than liver mirex concentrations in prey species (snowshoe

hare, Lepus americanus, 0.08 to 0.13 µg/kg FW; northern red-backed vole, Clethrionomys rutilus,

0.32 µg/kg FW), suggesting that mirex biomagnification in mammalian wildlife food chains ispossible (Poole et al 1998)

of mirex products should not exceed 232 µg per hour (USPHS 1995)

Before the banning of mirex for all uses in 1978, the tolerance limits in food for humanconsumption were 0.1 mg/kg for eggs, milk, and fat of meat from cattle, goats, hogs, horses, poultry,and sheep, and 0.01 mg/kg for all other raw agricultural commodities (Waters et al 1977; Buckler

et al 1981); higher limits of 0.3 mg mirex/kg in fish and shellfish and 0.4 mg/kg in crabs weretolerated (NAS 1978) The maximum recommended allowable concentration of mirex in edibleportions of domestic fish for human consumption was 0.1 mg/kg FW at that time (Scrudato andMcDowell 1989) Avoidance of larger and older fish to minimize ingestion of fat-soluble contam-inants, including mirex, was recommended Trimming the fatty tissues from muscle of salmon andtrout from Lake Ontario prior to consumption resulted in a mirex reduction of at least 44% in thetrimmed fillet — reflecting loss of fat content — and a product considered safe (i.e., <0.1 mgmirex/kg FW) by the U.S Food and Drug Administration (Insalaco et al 1982; Voiland et al 1991).However, mirex concentrations as low as 0.1 mg/kg in diets of adult prairie voles were associatedwith delayed maturation of pups, and with significant delays in the attainment of various earlydevelopment behaviors such as bar-holding ability, hind-limb placing, and negative geotaxis (Shan-non 1976) It is not known whether or not prairie voles can serve as a model for protection ofhealth of humans or various wildlife species In the absence of supporting data, however, it seemsprudent now to establish a dietary threshold of mirex at some level lower than 0.1 mg/kg Amaximum concentration of 0.01 mg/kg total dietary mirex, which is a recommended level for mostraw agricultural commodities, appears reasonable and conservative for the protection of fish,wildlife, and human health This value could be modified as new data become available.

Although mirex is extremely persistent in the environment, research findings suggest that somedegradation occurs and that some of the degradation products, such as photomirex, are biologicallyactive Accordingly, additional research is warranted on the fate and effects of mirex degradationproducts, with special emphasis on biomagnification through aquatic and terrestrial food chains.Alternate means of controlling imported fire ants are under consideration One approach hasbeen to reduce the concentrations of active mirex in bait formulations from 0.3% to some lower,but effective, level Paton and Miller (1980) demonstrated that mirex baits containing 0.07% mirexwere effective in controlling Australian termites, reporting a 90% kill in 9 days Baits containing

as little as 0.01% mirex were also reported effective, although termite mortality was delayedconsiderably Waters et al (1977) indicated that alternate chemical control agents, such as chlo-rpyrifos, diazinon, dimethoate, or methyl bromide may be suitable and that nonbiocidal chemicals,such as various pheromones and hormones, which are capable of disrupting reproductive behavior

of fire ants, are also under active consideration Another proposal was to chemically modify mirex

to a more water-soluble and rapidly degradable product (Waters et al 1977) The formulationFerriamicide, which consisted of 0.05% mirex, ferrous chloride, and a small amount of long-chainalkyl amines, was formulated in baits during 1978/79 for ant control (Lowe 1982) Ferriamicidedegraded within a few days after initial application; however, approval was revoked in 1980 when

it was learned that the toxicity of various degradation products to mammals, especially that ofphotomirex, exceeded that of 4X bait formulations (Lowe 1982)

Mirex replacements should not manifest the properties that led to the discontinuance of mirexfor all uses, namely:

• Delayed mortality in aquatic and terrestrial fauna

• Numerous birth defects

• Tumor formation

• Histopathology

• Adverse effects on reproduction, early growth, and development

• High biomagnification and persistence

• Disrupted energy metabolism

• Degradation into toxic metabolites

• Population alterations

• Movement through aquatic and terrestrial environmental compartments.

It is emphasized that mirex replacement compounds must be thoroughly tested before widespreadapplication in the environment; if testing is incomplete, it is almost certain that the nation’s fish

and wildlife resources will be adversely affected.In 1980, the use of Amdro thyl-2) (Lh)-pyrimidine) was conditionally approved by the U.S Environmental Protection Agency(Lowe 1982) Amdro reportedly has good ant control properties, degrades rapidly in sunlight, has

(tetrahydro-5,5-dime-a biologic(tetrahydro-5,5-dime-al h(tetrahydro-5,5-dime-alf life of less th(tetrahydro-5,5-dime-an 24 h, is nonmut(tetrahydro-5,5-dime-agenic, (tetrahydro-5,5-dime-and is rel(tetrahydro-5,5-dime-atively nontoxic to other th(tetrahydro-5,5-dime-antargeted species, except fish Amdro was more acutely toxic than mirex to fish

Mirex (dodecachlorooctahydro-1,3,4-metheno-2H-cyclobuta [c,d] pentalene) has been used extensively in pesticidal formulations to control the red imported fire ant (Solenopsis invicta), and

as a flame retardant in electronic components, plastics, and fabrics One environmental consequence

of mirex was the severe damage recorded to fish and wildlife in nine southeastern states and theGreat Lakes, especially Lake Ontario In 1978, the U.S Environmental Protection Agency bannedall further use of mirex, partly because of the hazards it imposed on nontarget biota These included:

• Delayed mortality and numerous birth defects in aquatic and terrestrial fauna

• Tumor formation

• Histopathology

• Wildlife population alterations

• Adverse effects on reproduction, early growth, and development

• High biomagnification and persistence

• Degradation into toxic metabolites

• Movement through aquatic and terrestrial environmental compartments

• Disrupted mammalian energy metabolism

• Detection of residues in human milk and adipose tissues

Among susceptible species of aquatic organisms, significant damage effects were recordedwhen concentrations of mirex in water ranged from 2 to 3 µg/L The recommended concentration

of 0.001 µg mirex/L affords an unusual degree of protection Evidence suggests that sensitivespecies of wildlife are adversely affected at 0.1 mg/kg of dietary mirex For comparison, tolerancelimits for mirex in food for human consumption range from 0.01 mg/kg for raw agriculturalcommodities to 0.1 mg/kg for eggs, milk, animal fat, and various seafood products Additionalresearch is needed on the fate of mirex degradation products and their effects on natural resources.Further, it is strongly recommended that environmental use of all mirex replacement compounds

be preceded by intensive ecological and toxicological evaluation

Ahrens, F.A., D.C Dyer, and W.E Lloyd 1980 Mirex kinetics in chickens Jour Toxicol Environ Health

6:835-842.

Baker, M.F 1964 Studies on possible effects of mirex bait on the bobwhite quail and birds Proc Annu Conf.

Southeast Assoc Fish Game Comm 18:153-159.

Barbehenn, K.R and W.L Reichel 1981 Organochlorine concentrations in bald eagles: brain/body lipid

relations and hazard evaluation Jour Toxicol Environ Health 8:325-330.

Barber, R.T and S.M Warlen 1979 Organochlorine insecticide residues in deep sea fish from 2500 m in the

Atlantic Ocean Environ Sci Technol 13:1146-1148.

Bell, M.A., R.A Ewing, and G.A Lutz 1978 Reviews of the Environmental Effects of Pollutants: I Mirex and Kepone U.S Environ Protection Agency Rep 600/1-78-013 235 pp.

Bird, D.M., P.H Tucker, G.A Fox, and P.C Lague 1983 Synergistic effects of Aroclor ® 1254 and mirex on

the semen characteristics of American kestrels Arch Environ Contam Toxicol 12:633-640.

Blumton, A.K., J.D Fraser, R.W Young, S Goodbred, S.L Porter, and D.L Luukkonen 1990 Pesticide and

PCB residues for loggerhead shrikes in the Shenandoah Valley, Virginia, 1985–88 Bull Environ Contam.

Toxicol 45:697-702.

Blus, L.J 1995 Organochlorine pesticides Pages 275-300 in D.J Hoffman, B.A Rattner, G.A Burton, Jr.,

and J Cairns, Jr (eds.) Handbook of Ecotoxicology Lewis Publishers, Boca Raton, FL.

Borgmann, U and D.M Whittle 1991 Contaminant concentration in Lake Ontario lake trout (Salvelinus

namaycush): 1977 to 1988 Jour Great Lakes Res 17:368-381.

Buckler, D.R., A Witt, Jr., F.L Mayer, and J.N Huckins 1981 Acute and chronic effects of kepone and

mirex on the fathead minnow Trans Amer Fish Soc 110:270-280.

Bush, B., J Snow, S Connor, L Rueckeit, Y Han, P Dymerski, and D Hilker 1983 Mirex in human milk

in upstate New York Arch Environ Contam Toxicol 12:739-746.

Cain, B.W and C.M Bunck 1983 Residues of organochlorine compounds in starlings (Sturnus vulgaris),

1979 Environ Monitor Assess 3:161-172.

Chernoff, N., R.E Linder, T.M Scotti, E.H Rogers, B.D Carver, and R Kavlock 1979 Fetotoxicity and

cataractogenicity of mirex in rats and mice with notes on kepone Environ Res 18:257-269.

Chu, I., D.C Villaneuve, V.E Secours, V.E Valli, and G.C Becking 1981 Effects of photomirex and mirex

on reproduction in the rat Toxicol Appl Pharmacol 60:549-556.

Clark, D.R., Jr and A.J Krynitsky 1980 Organochlorine residues in eggs of loggerhead and green sea turtles

nesting at Merritt Island, Florida — July and August 1976 Pestic Monitor Jour 14:7-10.

Clark, K.E and D Mackay 1991 Dietary uptake and biomagnification of four chlorinated hydrocarbons by

guppies Environ Toxicol Chem 10:1205-1217.

Cripe, C.R and R.J Livingston 1977 Dynamics of mirex and its principal photoproducts in a simulated

marsh system Arch Environ Contam Toxicol 5:295-303.

Custer, T.W., C.M Bunck, and T.E Kaiser 1983 Organochlorine residues in Atlantic coast black-crowned

night-heron eggs, 1979 Colonial Waterbirds 6:160-167.

de la Cruz, A.A and K.Y Lue 1978a Mirex incorporation in estuarine animals, sediment, and water,

Mississippi Gulf Coast — 1972–74 Pestic Monitor Jour 12:40-42.

de la Cruz, A.A and K.Y Lue 1978b Mirex incorporation in the environment: in situ decomposition of fire ant bait and its effects on two soil macroarthropods Arch Environ Contam Toxicol 7:47-61.

De Smet, K.D 1987 Organochlorines, predators and reproductive success of the red-necked grebe in southern

Manitoba Condor 89:460-467.

Dutil, J.D., B Legare, and C Desjardins 1985 Discrimination d’un stock de poisson, l’anguille (Anguilla

rostrata), basee sur la presence d’un produit chimique de synthese, le mirex Canad Jour Fish Aquatic Sci 42:455-458.

Eisler, R 1985 Mirex Hazards to Fish, Wildlife, and Invertebrates: A Synoptic Review U.S Fish Wildl Serv Biol Rep 85 (1.1) 42 pp.

Ervin, M.G 1982 The role of the Pituitary-Adrenal Cortical-Liver Axis in Mirex-Induced Adaptive Liver Hypertrophy Ph.D Thesis, Mississippi State Univ., Mississippi State 84 pp.

Eversole, A.G 1980 Effects of Water-Borne Mirex on the Survival and Production of Macrobrachium

rosen-bergii (de Man) Water Resour Res Inst., Clemson Univ., Clemson SC, OWRT Proj B-115-SC 44 pp.

Ewins, D.J., D.V Weseloh, and P Mineau 1992 Geographical distribution of contaminants and productivity

measures of herring gulls in the Great Lakes: Lake Huron 1980 Jour Great Lakes Res 18:316-330.

Ewins, P.J., D.V Weseloh, R.J Norstrom, K Lagierse, H.J Auman, and J.P Ludwig 1994 Caspian Terns on the Great Lakes: Organochlorine Contamination, Reproduction, Diet, and Population Changes, 1972–1991 Canad Wildl Serv., Occas Paper Numb 85 28 pp.

Farr, D.H and L.M Blake 1979 A creel census of the salmonid fishery in South Sandy Creek, New York.

N.Y Fish Game Jour 26:1-10.

Fujimori, K., I.K Ho, H.M Mehendale, and D.C Villeneuve 1983 Comparative toxicology of mirex,

photomirex and chlordecone after oral administration to the mouse Environ Toxicol Chem 2:49-60 Gaines, T.B and R.D Kimbrough 1969 The oral toxicity of mirex in adult and suckling rats Toxicol Appl.

Pharmacol 14:631-632.

Gilman, A.P., G.A Fox, D.B Peakall, S.M Teeple, T.R Carroll, and G.T Haymes 1977 Reproductive

parameters and egg contaminant levels of Great Lakes herring gulls Jour Wildl Manage 41:458-468.

Gilman, A.P., D.J Hallett, G.A Fox, L.J Allan, W.J Learning, and D.B Peakall 1978 Effects of injected

organochlorines on naturally incubated herring gull eggs Jour Wildl Manage 42:484-493.

Gobas, F.A.P.C., K.E Clark, W Y Shiu, and D Mackay 1989 Bioconcentration of polybrominated benzenes and biphenyls and related superhydrophobic chemicals in fish: role of bioavailability and elimination into

the feces Environ Toxicol Chem 8:231-245.

Gobas, F.A.P.C., X Zhang, and R Wells 1993 Gastrointestinal magnification: the mechanism of

biomagni-fication and food chain accumulation of organic chemicals Environ Sci Technol 27:2855-2863.

Grabowski, C.T 1981 Plasma proteins and colloid osmotic pressure of blood of rat fetuses prenatally exposed

to mirex Jour Toxicol Environ Health 7:705-714.

Halfon, E 1984 Error analysis and simulation of mirex behaviour in Lake Ontario Ecolog Model 22:213-252.

Halfon, E 1987 Modeling of mirex loadings to the bottom sediments of Lake Ontario within the Niagara

River plume Jour Great Lakes Res 13:18-23.

Hall, R.J., T.E Kaiser, W.B Robertson, Jr., and P.C Patty 1979 Organochlorine residues in eggs of the

endangered American crocodile (Crocodylus acutus) Bull Environ Contam Toxicol 23:87-90.

Heath, R.G., J.W Spann, E.F Hill, and J.F Kreitzer 1972 Comparative Dietary Toxicities of Pesticides to Birds U.S Fish Wildl Serv Spec Sci Rep — Wildl 152 57 pp.

Hill, E.P and D.M Dent 1985 Mirex residues in seven groups of aquatic and terrestrial mammals Arch.

Environ Contam Toxicol 14:7-12.

Holcombe, C.M and W.S Parker 1979 Mirex residues in eggs and livers of two long-lived reptiles (Chrysemys

scripta and Terrapene carolina) Bull Environ Contam Toxicol 23:369-371.

Hollister, T.A., G.E Walsh, and J Forester 1975 Mirex and marine unicellular algae: accumulation, population

growth and oxygen evaluation Bull Environ Contam Toxicol 14:753-759.

Holloman, M.E., B.R Layton, M.V Kennedy, and C.R Swanson 1975 Identification of the major thermal

degradation products of the insecticide mirex Jour Agric Food Chem 23:1011-1012.

Howard, F.W and A.D Oliver 1978 Arthropod populations in permanent pastures treated and untreated with

mirex for red imported fire ant control Environ Entomol 7:901-903.

Huckins, J.N., D.L Stalling, J.D Petty, D.R Buckler, and B.T Johnson 1982 Fate of kepone and mirex in

the aquatic environment Jour Agric Food Chem 30:1020-1027.

Hunter, R.G., J.H Carroll, and J.C Randolph 1980 Organochlorine residues in fish of Lake Texoma, October

1979 Pestic Monitor Jour 14:102-107.

Hyde, K.M 1972 Studies of the Responses of Selected Wildlife Species to Mirex Bait Exposure Ph.D Thesis, Louisiana State University, Baton Rouge 170 pp.

Innes, J.R.M., B.M Ulland, M.G Valerio, L Petrucelli, L Fishbein, E.R Hart, A.J Pallotta, P.R Bates, H.L Falk, J.J Gart, M Klein, I Mitchell, and J Peters 1969 Bioassay of pesticides and industrial chemicals

for tumorigenicity in mice: a preliminary report Jour Nat Cancer Inst 42:1101-1114.

Insalaco, S.E., J.C Makarewicz, and J.N McNamara 1982 The influence of sex, size, and season on mirex

levels within selected tissues of Lake Ontario salmon Jour Great Lakes Res 8:660-665.

Johnson, W.W and M.T Finley 1980 Handbook of Acute Toxicity of Chemicals to Fish and Aquatic

Inver-tebrates Summaries of Toxicity Tests Conducted at Columbia National Fisheries Research Laboratory,

1965–78 U.S Fish Wildl Serv Resour Publ 137 98 pp.

Jones, A.S and C.S Hodges 1974 Persistence of mirex and its effects on soil microorganisms Jour Agric.

Food Chem 22:435-439.

Kaiser, K.L.E 1978 The rise and fall of mirex Environ Sci Technol 12:520-528.

Kavlock, R.J., N Chernoff, E Rogers, D Whitehouse, B Carver, J Gray, and K Robinson 1982 An analysis

of fetotoxicity using biochemical endpoints of organ differentiation Teratology 26:183-194.

Kendall, R.J., R Noblet, J.D Hair, and H.B Jackson 1977 Mirex residues in bobwhite quail after aerial

application of bait for fire ant control, South Carolina — 1975–76 Pestic Monitor Jour 11:64-68.

Kendall, R.J., R Noblet, L.H Senn, and J.R Holman 1978 Toxicological studies with mirex in bobwhite

quail Poult Sci 57:1539-1545.

Kim, H.T., K.S Kim, J.S Kim, and W.B Stone 1985 Levels of polychlorinated biphenyls (PCBs), DDE,

and mirex in waterfowl collected in New York state, 1981–82 Arch Environ Contam Toxicol 14:13-18.

Kim, K.S., M.J Pastel, J.S Kim, and W.B Stone 1984 Levels of polychlorinated biphenyls, DDE, and mirex

in waterfowl collected in New York state, 1979–80 Arch Environ Contam Toxicol 13:373-381.

Klaas, E.E., H.M Ohlendorf, and E Cromartie 1980 Organochlorine residues and shell thicknesses in eggs

of the clapper rail, common gallinule, purple gallinule, and limpkin (Class Aves), Eastern and Southern

United States, 1972–74 Pestic Monitor Jour 14:90-94.

Kobylinski, G.J and R.J Livingston 1975 Movement of mirex from sediment and uptake by the hogchoker,

Trinectes maculatus Bull Environ Contam Toxicol 14:692-698.

Koenig, C.C 1977 The effects of DDT and mirex alone and in combination on the reproduction of a salt

marsh cyprinodont fish, Adinia xenica Pages 357-376 in F.J Vernberg, A Calabrese, F.P Thurberg, and W.B Vernberg (eds.) Physiological Responses of Marine Biota to Pollutants Academic Press, New York.

Larson, P.S., J.L Egle, Jr., G.R Hennigar, and J.F Borzelleca 1979 Acute and subchronic toxicity of mirex

in the rat, dog, and rabbit Toxicol Appl Pharmacol 49:271-277.

Leatherland, J.F and R.A Sonstegard 1981 Effect of dietary mirex and PCB’s on calcium and magnesium

metabolism in rainbow trout, Salmo gairdneri and coho salmon, Oncorhynchus kisutch; a comparison with Great Lakes coho salmon Comp Biochem Physiol 69C:345-351.

Leatherland, J.F., R.A Sonstegard, and M.V Holdrient 1979 Effect of dietary mirex and PCB’s on somatic index, liver lipid, carcass lipid and PCB and mirex bioaccumulation in yearling coho salmon,

hepato-Oncorhynchus kisutch Comp Biochem Physiol 63C:243-246.

Lee, J.H., C.E Nash, and J.R Sylvester 1975 Effects of Mirex and Methoxychlor on Striped Mullet, Mugil

cephalus L.U.S Environ Protection Agency Rep EPA-660/3-75-015 18 pp.

Lewis, T.W and J.C Makarewicz 1988 Exchange of mirex between Lake Ontario and its tributaries Jour.

Great Lakes Res 14:388-393.

Lowe, J.I 1982 Mirex, fire ants, and estuaries Pages 63-70 in Proceedings of the Workshop on Agrichemicals

and Estuarine Productivity Duke Univ Mar Lab., Beaufort, NC Sept 18-19, 1980 U.S Dep Comm.

NOAA/OMPA.

Ludke, J.L., M.T Finley, and C Lusk 1971 Toxicity of mirex to crayfish, Procambarus blandingi Bull.

Environ Contam Toxicol 6:89-96.

Lue, K.Y and A de la Cruz 1978 Mirex incorporation in the environment: toxicity in Hydra Bull Environ.

Contam Toxicol 19:412-416.

Markin, G.P 1981 Translocation and fate of the insecticide mirex within a bahia grass pasture ecosystem.

Environ Pollut 26A:227-241.

McArthur, M.L.B., G.A Fox, D.B Peakall, and B.J.R Philogene 1983 Ecological significance of behavioral

and hormonal abnormalities in breeding ring doves fed an organochlorine mixture Arch Environ Contam.

Toxicol 12:343-353.

McCorkle, F.M., J.E Chambers, and J.D Yarbrough 1979 Tissue enzyme activities following exposure to

dietary mirex in the channel catfish, Ictalurus punctatus Environ Pollut 19:195-199.

McLane, M.A.R., E.H Dustman, E.R Clark, and D.L Hughes 1978 Organochlorine insecticide and

poly-chlorinated biphenyl residues in woodcock wings, 1971–72 Pestic Monitor Jour 12:22-25.

Mehrle, P.M., F.L Mayer, and D.R Buckler 1981 Kepone and mirex: effects on bone development and swim

bladder composition in fathead minnows Trans Amer Fish Soc 110:638-643.

Menzie, C.M 1978 Metabolism of Pesticides Update II U.S Fish Wildl Serv Spec Sci Rep — Wildl.

212 381 pp.

Minchew, C.D., R.N Hunsinger, and R.C Giles 1980 Tissue distribution of mirex in adult crayfish

(Pro-cambarus clarki) Bull Environ Contam Toxicol 24:522-526.

Muncy, R.J and A.D Oliver 1963 Toxicity of ten insecticides to the red crawfish, Procambarus clarki (Girard) Trans Amer Fish Soc 92: 428-431.

Naber, E.C and G.W Ware 1965 Effects of kepone and mirex on reproductive performance in the laying

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Board Canada 35:1401-1409.

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Lakes herring gulls Environ Sci Technol 14:860-866.

Ohlendorf, H.M., J.C Bartonek, G.J Divoky, E.E Klaas, and A.J Krynitsky 1982 Organochlorine Residues

in Eggs of Alaskan Seabirds U.S Fish Wildl Serv Spec Sci Rep — Wildl 245 1-41.

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Pestic Monitor Jour 14:125-135.

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Toxicol 12:355-382.

Schoor, W.P 1979 Distribution of mirex in an experimental estuarine ecosystem Bull Environ Contam.

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Schoor, W.P and S.M Newman 1976 The effect of mirex on the burrowing activity of the lugworm (Arenicola

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Shannon, V.C 1976 The Effects of Mirex on the Reproductive Performance and Behavioral Development of

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Skaar, D.R., B.T Johnson, J.R Jones, and J.N Huckins 1981 Fate of kepone and mirex in a model aquatic

environment: sediment, fish, and diet Canad Jour Fish Aquat Sci 38:931-938.

Skea, J.C., H.J Simonin, S Jackling, and J Symula 1981 Accumulation and retention of mirex by brook

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Environ Sci Technol 29:740-750.

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9:73-78.

CHAPTER 22 Paraquat

Paraquat (1,1′-dimethyl-4,4′-bipyridinium) is one of the most widely used herbicidal chemicals

in the world and is now available in more than 130 countries (Kimbrough 1974; Calderbank 1975;Dasta 1978; Haley 1979; Hughes 1988; Smith 1988b; Eisler 1990) Its chemical structure was firstdescribed in 1882, its oxidizing and reducing properties in 1933, and its herbicidal properties in 1955.Paraquat was marketed commercially in the United Kingdom in 1962 and registered for use in theUnited States in 1964 As the dichloride salt, it has found wide use as a nonselective contact herbicide

at application rates of 1.12 kg/ha (1 pound/acre) and lower Paraquat kills plants by affecting the greenparts, not the woody stems, and is usually completely and rapidly inactivated by contact with clay inthe soil In its bound form, paraquat is biologically inert and innocuous to plants and animals (Fletcher1974) In 1977, the discovery by narcotics authorities that some marijuana imported from Mexicohad been treated with paraquat as a control agent generated much interest in the media (Dasta 1978;Haley 1979) Up to 70% of the paraquat in paraquat-treated marijuana, on smoking, is converted tobipyridine, a respiratory irritant Frequent consumption of heavily contaminated cigarettes may result

in cyanosis and possibly death The use of paraquat for this purpose has been largely discontinued.Numerous human injuries and deaths have resulted from intentional ingestion of the concentratedcommercial product (Fletcher 1974; Dasta 1978; Haley 1979; Crome 1986; Smith 1988a, 1988b).For example, in the first 10 years following paraquat commercial use, 232 human deaths fromparaquat poisoning were reported, about half suicidal, and almost all were due to the drinking ofconcentrated material Most poisonings resulted from the ingestion of the 21% cation concentrate,which had been decanted and stored in empty beer, soft drink, or lemonade bottles; paraquat is areddish-brown liquid that resembles root beer or cola drinks One individual sprinkled paraquat onFrench fried potatoes, thinking that it was vinegar He died 25 days later Another died after applyingthe concentrated solution to his beard and scalp to treat a lice infestation In Japan, more than

1000 persons each year are reportedly poisoned by paraquat Initially, paraquat may produce tiorgan toxicity of kidneys, liver, heart, central nervous system, adrenal glands, skeletal muscle, andspleen, but the ultimate target organ is the lung, in which progressive irreversible pulmonary fibrosisdevelops This effect has been described in man, rats, mice, guinea pigs, and dogs (Kimbrough 1974;Giri et al 1979; Hampson and Pond 1988; O’Sullivan 1989) At present, there is no specific antidotefor paraquat poisoning In normal use as a spray, minor reversible injuries are reported to abradedskin, eyes, nose, and fingernails; it is not absorbed through intact skin (Kimbrough 1974; Smith1988a) Paraquat is fetotoxic as judged by deliberate ingestion of concentrated solutions by ninepregnant Taiwanese women Paraquat crosses the placenta and concentrates there to levels 4 to 6times that of maternal blood All fetuses died whether or not an emergency cesarean operation wasperformed (Talbot and Fu 1988) Research has focused on the tendency of paraquat to accumulate

mul-in neuromelanmul-in of mammals and amphibians and to cause lesions mul-in the pigmented nerve cells,leading to effects very similar to those of Parkinson’s disease (De Gori et al 1988; Lindquist et al.1988) Reviews on ecological and toxicological aspects of paraquat include those by Kimbrough(1974), Smith and Heath (1976), Autor (1977), Dasta (1978), Haley (1979), Summers (1980), Bauer(1983), Onyeama and Oehme (1984), Smith (1985, 1988a, 1988b), and Eisler (1990).

Paraquat is a broad-spectrum contact weed killer and herbage desiccant that is used widely inagriculture and horticulture Paraquat was originally formulated in 1882, but its herbicidal propertieswere not discovered until 1955 Since its introduction in the early 1960s, paraquat has been usedextensively in about 130 countries, including the United Kingdom, Canada, and the United States, on

a wide variety of agricultural crops (Fletcher 1974; Haley 1979; Kelly et al 1979; Anonymous 1988).Primary uses of paraquat include: weed control in orchards, plantation crops, and forests; weedcontrol before sowing or before crop emergence; pasture renovation; preharvest desiccation; andaquatic weed control, although use as an aquatic herbicide in the United States is not permitted(Anonymous 1963, 1974; Summers 1980; Dial and Bauer 1984) In New Zealand, use of paraquatfor aquatic weed control (2 mg/L for 30 min) in 1966/67 on the Waimakariri River severely reducedamphipod populations; paraquat is no longer used for this purpose in that country (Burnet 1972).Paraquat is registered domestically for preplant or preemergence use for cotton, barley, corn, lettuce,melons, peppers, safflower, soybeans, sorghum, sugar beets, tomatoes, potatoes, and wheat It isalso registered for use on noncrop areas, such as roadsides, highway margins, rights-of-way, aroundcommercial buildings, power plants, storage yards, fence lines, and parkways (Anonymous 1963,1974) In Switzerland, it is used to control voles (Arvicola terrestris) in fruit orchards (Summers1980) Paraquat application to corn using a manual knapsack sprayer is considered unsafe to thehuman operators if lances are less than 1.0 m Lances >1.0 m in length are recommended, as well

as additional protective garments for legs, feet, and hands and switching the paraquat sprayingoperation to the back of the worker’s body (Machado-Neto et al 1998)

Paraquat is available as the dichloride or dimethylsulfate salt; both compounds are extremelysoluble in water (Kimbrough 1974) In the United States, paraquat dichloride is available as a 29%liquid concentrate containing 240 g/L (2 pounds/gallon) of paraquat cation, or as a 42% liquidconcentrate Elsewhere, it is sold as Gramoxone liquid containing 20 to 24% paraquat dichloride(Fletcher 1974; Bauer 1983; Dial and Dial 1987a) Paraquat dichloride concentrates usually containvarious wetting agents (condensation products of ethylene oxide and alkyl phenols), spreaders, humec-tants to promote moisture retention (calcium chloride, glycerol, polyethylene glycol), plant adhesionmaterials (carboxymethylcellulose, polymethacrylates), and antifoaming agents (Summers 1980).The recommended field application rates for terrestrial weed control usually range between0.28 and 1.12 kg paraquat cation/ha (0.25 and 1.0 pounds/acre), between 0.56 and 2.24 kg paraquatdichloride/ha (0.5 and 2.0 pounds/acre) — both applied as an aerosol — and between 0.1 and2.0 mg/L for aquatic weed control, although sensitive aquatic plants may be affected between0.019 and 0.372 mg/L (Ross et al 1979; Summers 1980; Bauer 1983; Dial and Bauer 1984).Paraquat is frequently used in combination with other herbicides (Fletcher 1974; Summers 1980).Water solutions of the dichloride salt, which usually contain 240 g/L, have been successfully mixedwith 2,4-D, substituted ureas, dalapon, amitrol, and various triazines (Anonymous 1963, 1974)

Data are scarce on ecosystems treated with paraquat It is clear, however, that both terrestrialand aquatic plants accumulate paraquat, and that the compound disappears rapidly from the watercolumn and tends to concentrate in surface muds (Table 22.1)

Water from irrigation channels, rivers, and lagoons from Spanish marshes in 1996 borderingthe Mediterranean Sea contained an average of 0.01 µg paraquat/L, with a maximum recordedvalue of 3.95 µg/L (Fernandez et al 1998) Paraquat values were highest in the summer owing tohigh application rates, low rainfall, and high evaporation rates At these comparatively low con-centrations, paraquat is not easily degraded chemically or biologically and persists in river waterswith more than 80% remaining after 56 days of incubation (Fernandez et al 1998).

Table 22.1 Paraquat Concentrations in Field Collections of Selected Organisms

and Nonbiological Materials

Sample and Other Variables

Concentration (mg/kg dry weight) Reference a TREATED FIELDS

death, are not unusual No treatment or chemical has proven completely successful in protectingagainst paraquat-induced lung toxicity.

Paraquat is strongly adsorbed to soils and sediments and is biologically unavailable in thatform; however, it is not degraded significantly for many years, except in surface soils In surfacesoils, paraquat loss through photodecomposition approaches 50% in 3 weeks In freshwater eco-systems, loss from water column is rapid: about 50% in 36 h and 100% in 4 weeks In marineecosystems, 50 to 70% loss of paraquat from seawater was usually recorded within 24 h

22.4.2 Chemical Properties

Paraquat is a nonvolatile, ionic compound that is almost completely insoluble in fat, and thereforenot likely to be accumulated in food chains (Calderbank 1975) The compound belongs to thebipyridyl group of chemicals and is typical of the many hundreds that have been synthesized, variationusually being the result of introducing different quaternizing groups on the nitrogen atoms, whichalso shift (Fletcher 1974) (Table 22.2) Paraquat dichloride is produced from pyridine in the presence

of sodium in anhydrous ammonia, then quaternizing the 4,4′-dipyridyl with methyl chloride (Haley1979) The common paraquat salts are all fully ionized, and experiments have shown that the anions(e.g., chloride, sulfate, methyl sulfate) do not affect the toxicity of paraquat (Fletcher 1974) Chemicaland other properties of paraquat are briefly summarized in Figure 22.1 and Table 22.2

22.4.3 Mode of Action

Paraquat is absorbed systematically in mammals, following different routes of exposure; tion is greatest for the pulmonary route, followed by intragastric and dermal routes (Chui et al.1988) Administration of paraquat by every route of entry tested frequently results in irreversiblechanges in lung (Boudreau and Nadeau 1987) In the intestinal tract, where some microbialdegradation occurs, most paraquat (95 to 100%) is usually excreted unchanged in feces and urinewithin 2 days (Summers 1980) Absorption in the gastrointestinal tract ranges from 0.26% in cow,

absorp-to 5% in man, 8% in guinea pig, 16% in cat, and up absorp-to 20% in rat The half-time persistence(Tb 1/2) of paraquat in certain tissues ranges between 20 and 30 min, but up to 4 days in muscleand 2 days in plasma (Bauer 1983) Delayed toxic effects of paraquat occurring after the excretion

of virtually all of the material have caused it to be classified as a “hit and run” compound, that is,

a compound causing immediate damage, the consequences of which are not readily apparent(Conning et al 1969)

Most authorities agree that free radical pathology is the most likely mechanism by whichparaquat is cytotoxic (Bus et al 1976; Frank et al 1982; Patterson and Rhodes 1982; Combs andPeterson 1983; Onyeama and Oehme 1984; Gabryelak and Klekot 1985; Smith 1985; Wong andStevens 1986; Seto and Shinohara 1987; Suleiman and Stevens 1987; Darr et al 1988; Dunbar

et al 1988a; Wegener et al 1988; Wenning et al 1988; Arias et al 1991; Babich et al 1993) Thebiochemical mechanism of paraquat toxicity is related to the cyclic oxidation and reduction ofparaquat that occurs in lung cells, which leads to continued production of high levels of superoxide

Figure 22.1 Structural formulas of paraquat cation

(upper) and paraquat dichloride salt (lower).

anion (O2) and other cytotoxic oxygen free radicals Superoxide anion and other oxygen freeradicals initiate the peroxidation of membrane lipids, causing tissue damage and death Paraquatoxidation is coupled with the reduction of molecular oxygen, forming superoxide anion, singletoxygen, and hydroxyl radicals These molecular species react with polyunsaturated fatty acid freeradicals and, on further oxidation, with lipid hydroperoxide radicals The hydroperoxide radicalsthen maintain the formation of new fatty acid radicals while being converted to lipid peroxides in

a chain reaction Various enzymes in the cells catabolize the superoxide radical and reduce the lipidhydroperoxides to less-toxic lipid alcohols The superoxide anions are converted to hydrogenperoxide and oxygen; hydrogen peroxide is further inactivated to water and oxygen by catalasesand peroxidases In the presence of reduced nicotinamide adenine dinucleotide phosphate(NADPH), paraquat is reduced by microsomal NADPH-cytochrome reductase The reduction oflipid peroxides by glutathione peroxidase requires reduced glutathione Because the reduction ofoxidized glutathione is coupled with NADPH oxidation via glutathione reductase, it seems that theavailability of NADPH is essential for paraquat detoxification, and that the critical depletion ofNADPH may render the cell more susceptible to lipid peroxidation

The lung is the organ most severely affected in paraquat poisoning (Campbell 1968; Conning

et al 1969; Haley 1979; Aldrich et al 1983; Bauer 1983; Combs and Peterson 1983; Christian

et al 1985; Smith 1985; Wong and Stevens 1986; Boudreau and Nadeau 1987; Baud et al 1988;Dunbar et al 1988a; Wegener et al 1988) Pulmonary injury is due largely to the preferentialaccumulation of paraquat in lung — mediated by an energy-dependent system for uptake ofendogenous polyamines — and to the continuous exposure of lung to atmospheric oxygen Char-acteristic signs of poisoning include severe anoxia, marked and widespread fibroblastic proliferation

in the alveolar walls around the terminal bronchi and blood vessels, and frequently death Thespecific toxicity to the lung can be explained by the accumulation of paraquat in the alveolar Type IIcells These cells are responsible for the synthesis of pulmonary surfactant, the surface-activematerial lining the alveolar epithelium The pulmonary surfactant is secreted after storage in

Table 22.2 Chemical and Other Properties of Paraquat

Variable Datum

Chemical name

Paraquat (cation) 1,1 ′ -dimethyl-4,4 ′ -bipyridinium

Paraquat dichloride (salt) 1,1 ′ -dimethyl-4,4 ′ -bipyridinium dichloride

Molecular weight 186.2 (cation), 257.2 (salt)

CAS Number 4685-14-17 (cation), 1910-42-5 (salt)

Empirical formula C12H14N2 (cation), C12H14Cl2N2 (salt)

Alternate names Cekuquat, Crisquat, Dextrone, Dextrone X, Dexuron, Dual Paraquat, Esgram,

Gramonol, Gramoxone, Gramuron, Herbaxon, Herboxone, Methyl Viologen, Ortho Paraquat, Orvar, Paracol, Paraquat CL, Pathclear, Pillarquat, Pillarxone, Preeglone, PP 148, PP 910, Sweep, Tenaklene, Totacol, Toxer Total, Weedol Solubility at 20°C

Most organic solvents Insoluble or sparingly soluble

Physical state White (pure), yellow (technical) solid

Main uses Herbicide, desiccant

Specific gravity 1.24–1.26

Melting point 175 – 180°C, decomposes at 345°C

Stability Stable on exposure to hot acids, unstable in alkalis at pH > 10

Flash point Nonexplosive, nonflammable

Data from Anonymous 1963, 1974, 1988; Haley 1979; Kelly et al 1979; Johnson and Finley 1980; Hudson et al 1984; Hill and Camardese 1986; Mayer 1987.

cytoplasmic organelles known as lamellar bodies Thus, any damage to the alveolar epitheliumcould alter synthesis and secretion of the pulmonary surfactant The pulmonary effects of paraquatare probably related to the conversion of paraquat to a free radical, followed by conversion to along-lived dihydroderivative, which causes transformation of normal alveolar epithelial cells tofibroblasts The increase in toxicity of paraquat by oxygen supports the hydroperoxide theory, inwhich the reversible action of the free radical’s oxidation-reduction gives rise to hydrogen peroxide.Paraquat also depletes NADPH in the isolated lung to the extent of mixed function oxidationimpairment Depletion of NADPH would impair fatty acid and lipoprotein synthesis and inhibitvarious detoxification and biosynthetic functions.

Other organs and systems affected by paraquat include the kidney (pathology of proximal tubules),liver (hepatocellular necrosis), spleen and thymus (pathology), circulatory system (irregular and feebleheart beat, myocardial congestion, increase in erythrocytes and leucocytes, external pericarditis,myocardium edema), brain (neuronal depletion, myelin destruction), gastrointestinal tract (esophagitis;ulceration of buccal cavity, pharynx, gastric mucosa; mucosal erosion), skin (erythema, hyperkerato-sis), reproductive system (degeneration), nervous system (hyperexcitability, irritability, incoordination,convulsions), various enzyme systems, and the eye (Giri et al 1979, 1982, 1983; Summers 1980;Bauer 1983; Seto and Shinohara 1987, 1988; Hughes 1988; Takegoshi et al 1988)

Several early indicators of paraquat-induced stress have been proposed, including alkalinephosphatase activity, fibronectin levels, and intracellular calcium uptake Alkaline phosphataseactivity is associated with the lamellar body, and changes in this variable are suggested as indicative

of toxicity to Type II alveolar epithelium cells (Boudreau and Nadeau1987) Levels of fibronectin,

an extracellular matrix glycoprotein, were elevated in patients with fibrotic lung diseases and inmonkeys given multiple injections of paraquat (Dubaybo et al 1987) Lung intracellular calciumuptake was significantly disrupted, even at doses that normally produce significant increases inlung water content (Agarwal and Coleman 1988) These subjects seem to merit additional research,

as does the role of polyamines in mediating fibrotic changes in the lung (Dunbar et al 1988b);paraquat-altered synthesis of proteins, DNA, collagen, and pentose phosphate metabolism (Simon

et al 1983); and hyperoxia, that is, increased oxygen-free radical generation (Frank et al 1982).Certain treatments or chemicals provide varying degrees of protection against paraquat-inducedlung toxicity and lethality, although no treatment or chemical has proven completely successful.Present treatment of paraquat-poisoned animals and humans is directed to elimination of the materialfrom the body using repeated doses of adsorbents such as Fuller’s earth or bentonite, cathartics toreduce paraquat absorption, and hemodialysis, forced diuresis, and hemofiltration to enhanceexcretion (Fletcher 1974; Autor 1977; Haley 1979; Pond et al 1987; Kitakouji et al 1989) Theuse of 100% oxygen is contraindicated, as mortality is greatly increased (Fletcher 1974; Autor1977; Wong and Stevens 1986) Toxicity mediated by free radicals can be moderated by severalcellular defense mechanisms, including superoxide dismutase, catalase, glutathione peroxidase,Vitamin E, and reduced glutathione (Gabryelak and Klekot 1985; Wenning et al 1988) A low-molecular-weight superoxide dismutase mimic, based on manganese, was found to protect mam-malian cells against the cytotoxic effects of the superoxide radical produced by paraquat (Darr

et al 1988) Certain chemicals reportedly provide limited protection to small laboratory animalsunder carefully controlled conditions of administration: nicotinic acid (Shibata and Iwai 1988);niacin (Heitkamp and Brown 1982); cysteine (Szabo et al 1986); N-acetylcysteine (Wegener et al.1988); metallothionein — a metal-binding, low-molecular-weight, protein rich in cysteine (Sato

et al 1989); d-penicillamine (Szabo et al 1986); clofibrate (Frank et al 1982); lipid-soluble oxidants (Kohen and Chevion 1988; Wegener et al 1988); various amino acids (Heitkamp andBrown 1982); phenobarbital (Bus et al 1976; Summers 1980); methyl prednisolone (Kitazawa et al.1988); and certain anti-inflammatory drugs (Autor 1977) In bluegills, 1,10-phenanthroline, achelator of ionic iron, reduced the toxicity of paraquat through the prevention of hydroxyl radicalformation (Babich et al 1993) In tilapia, the paraquat-induced increase in gill carbonic anhydraseactivity was not observed when ionic lead was present at 47 mg/L (Arias et al 1991) In plants,

anti-the pea (Pisum sativum) is protected by cerium chloride, in part, through counteracting peroxideformation (Vaughn and Duke 1983).

Paraquat toxicity is increased and its effects otherwise exacerbated in organisms fed dietsdeficient in selenium or Vitamin E, although high levels of these substances in diets did not provideprotection (Autor 1977; Haley 1979; Summers 1980); by methyl prostaglandins (Williams et al.1988) or diethyl maleate (Summers 1980); and by increased iron and copper (Kohen and Chevion1988; Ogino and Awai 1988) Dietary changes that do not result in nutrient deficiency or toxicitymay affect the biocidal properties of paraquat and other compounds In studies with rodentssubjected to paraquat insult, survival was higher in those fed cereal-based diets vs purified diets,and higher in egg-white (protein) purified diet vs a casein diet (Tanaka et al 1981; Evers et al.1982), suggesting a need to use strictly defined diets in the study of paraquat toxicity to controlfor any paraquat–diet interactions

Paraquat causes tissue damage and increased stress in the common carp (Cyprinus carpio), asjudged by the increased enzyme activities of lactic dehydrogenase, glutamic oxaloacetic transam-inase, and glutamate dehydrogenase, and by the elevated blood sugar levels Paraquat and coppersulfate when administered together to carp were synergistic in terms of tissue damage and stresseffects, especially liver damage (Asztalos et al 1990)

Paraquat adhering to the plant surface is usually degraded photochemically (Haley 1979;Summers 1980) Paraquat is phytotoxic through inhibition of processes involving photosynthesisand respiration (Haley 1979; Christian et al 1985; Anonymous 1988) Its mode of action in plants

is similar to that in animals; that is, lipid peroxidation of membranes due to formation of thesuperoxide radical and related species (Summers 1980) Photosynthetic tissues reduce paraquat tostable free radicals that, upon reoxidation, produce hydrogen peroxide Unsaturated lipids in thecells are oxidized by the peroxide, and damage is dependent largely on production of hydrogenperoxide (Haley 1979; Vaughn and Duke 1983) The reaction is light and oxygen dependent(Conning et al 1969; Kelly et al 1979)

In bacteria (Escherichia coli), paraquat is concentrated, reduced to the monocation radical, andcombines with molecular oxygen to produce the superoxide radical within the cell Copper andiron are essential mediators in bactericidal effects The cytoplasmic membrane is the target organelle

in paraquat toxicity to E coli, and extent of damage correlates positively with levels of these metals(Kohen and Chevion 1988)

22.4.4 Fate in Soils and Water

In contact with soil, paraquat is rapidly adsorbed — usually in the clay mineral lattice sheets —and inactivated by base exchange The process is facilitated by the flat and highly polarizable nature

of the paraquat ion (Anonymous 1963; Conning et al 1969; Calderbank 1975; Summers 1980;Kearney et al 1985) The strong binding of paraquat to soil constituents reduces the mobility of theherbicide due to leaching, although paraquat is displaced from binding sites by low concentrations

of ions of ammonium, potassium, sodium, and calcium (Smith and Mayfield 1978) Paraquat tion is not significantly affected by soil pH, but is modified by soil porosity, moisture content, residencetime, and adsorption capacity (Smith and Mayfield 1978; Summers 1980) Paraquat applied to a sandyloam soil at field application rates between 0.56 and 2.24 kg/ha was adsorbed by organic matter andclays, usually in the top centimeter of soil (Smith and Mayfield 1978) Typical soils contain about

adsorp-300 mg paraquat/kg after treatment at recommended applications; however, adsorption capacity variesamong soils Clay minerals, such as kaolinite, can adsorb 2500 to 3500 mg/kg, whereas others, such

as montmorillonite, adsorb up to 85,000 mg/kg after paraquat treatment (Summers 1980)

Paraquat is not degraded significantly in soil during incubation periods up to 16 months at 25°C

by chemical or microbiological vectors (Smith and Mayfield 1978) For example, paraquat ride applied once annually at 4.48 kg/ha, or 4 times annually at 1.12 kg/ha, remained essentiallyundegraded in the soil for 6 years (Fryer et al 1975; Moyer and Lindwall 1985) Massive applications

dichlo-to soils of 3000 kg/ha can persist for at least 6 months without significant degradation (Moyer andLindwall 1985) Bacterial degradation — which occurs only slowly in soils — consists of demethyla-tion, followed by ring cleavage to eventually form the carboxylated 1-methylpyridinium ion(Figure 22.2) Photochemical decomposition of paraquat is the predominant mechanism of paraquatdegradation in soils (Smith and Mayfield 1978) In surface soils, paraquat loss through photodecom-position was 20 to 50% in 3 weeks (Christian et al 1985) Photochemical degradation products ofparaquat include 4-carboxy-1-methylpyridium ion and methylamine hydrochloride (Figure 22.2) Lab-oratory studies have demonstrated that paraquat in soils slated for disposal can be degraded by ultraviolet(UV) irradiation in the presence of oxygen or ozone Reaction products identified were 4-carboxy-1-methylpyridium ion, 4-picolinic acid, hydroxy-4-picolinic acid, succinic acid, N-formylglycine, malicacid and oxalic acid (as trimethylsilicon derivatives), and 4,4′-bipyridyl (Kearney et al 1985).Paraquat is used to control aquatic weeds It also passes into aquatic environments through rain,where it is rapidly accumulated by aquatic organisms, especially fish (Gabryelak and Klekot 1985).Paraquat applied to control aquatic weeds is accumulated by aquatic macrophytes and algae, and

it is adsorbed to sediments and suspended materials Initial applications of 1 to 5 mg/L in the watercolumn are usually not detectable under field conditions after 8 to 27 days (Summers 1980) Thehalf-time persistence of paraquat in water column at normal doses for weed control (i.e., 0.5 to1.0 mg/L) was 36 h; less than 0.01 mg/L was detectable in 2 weeks (Calderbank 1975) In solution,paraquat was subject to photodecomposition and microbial metabolism, degrading to methylamine

Figure 22.2 Proposed pathway of paraquat degradation by a bacterial isolate (upper) and by ultraviolet

irradi-ation (lower) (Modified from Funderburk, H.H., Jr and G.A Bozarth 1967 Review of the olism and decomposition of diquat and paraquat Jour Agric Food Chem. 15:563-567.

metab-and 4-carboxy-1-methylpyridium ion (Kearney et al 1985) In freshwater, without sediment or plants,100% of the initial concentration of 0.5 mg paraquat/L was degraded in 35 weeks When sedimentswere present, 100% loss from the water column occurred in 6 to 8 weeks; and when both sedimentand aquatic plants were present, paraquat was not detectable in the water column in 3 to 4 weeks(Summers 1980) Mud cores taken from a paraquat-treated lake had elevated paraquat residues, butshowed no phytotoxic effects on barley seedlings germinated on them (Calderbank 1975).

Paraquat loss from seawater in 24 h was 70% at an initial concentration of 1 mg/L, 68% at

5 mg/L, and 76% at 10 mg/L; most of the loss occurred within the first 60 min (Fytizas 1980)

22.5.1 General

Adverse effects of paraquat in sensitive species of terrestrial plants and soil microflora have beendocumented at application rates of 0.28 to 0.6 kg/ha (death, inhibited germination of seeds, reducedgrowth), at soil concentrations of 10 to 25 mg/kg (growth inhibition), and at soil-water concentrations

as low as 1.6 mg/L (reduced growth, inhibited synthesis of protein and RNA) Among terrestrialinvertebrates, certain species of mites were sensitive to paraquat at recommended rates of application,and the sensitive honey bee died when its diet contained 100 mg/kg However, paraquat in soils wasnot accumulated by earthworms and other species of soil invertebrates after applications up to

112 kg/ha These points, and others listed in this section, are discussed in greater detail later.Freshwater algae and macrophytes usually die at paraquat concentrations between 0.25 and0.5 mg/L Marine algae, however, are relatively resistant and usually require 5 mg/L or higher forsignificant inhibition in growth to occur in 10 days Aquatic invertebrates, especially crustaceans,seem to be the most sensitive group, with effects most pronounced at elevated temperatures in earlydevelopmental stages Adverse effects were noted in crab larvae at nominal water concentrationsbetween 0.9 and 5.0 µg/L, although 1000 µg/L and higher were needed to produce similar effects

in other species of aquatic invertebrates Amphibians and fishes were usually unaffected at centrations below 3000 µg/L, although sensitive species, such as frog tadpoles and northern carp(Cyprinus carpio), were impacted at 500 µg/L There was little accumulation of paraquat from themedium by aquatic fauna

con-Paraquat is embryotoxic to sensitive species of birds Concentrations equivalent to 0.056 kg/haapplied in oil solution to the surface of eggs of the mallard (Anas platyrhynchos) inhibited devel-opment; when applied in aqueous solution, paraquat was toxic at a dose equivalent to 0.56 kg/ha

In each case, adverse effects occurred below the recommended field application rate of about1.0 kg/ha The lowest doses of paraquat that produced harmful effects in sensitive birds were

10 mg/kg body weight (BW) in nestlings of the American kestrel (Falco sparverius), 20 mg/kg inthe diet of northern bobwhite (Colinus virginianus), 40 mg/L in the drinking water of domesticchickens (Gallus sp.), and 199 mg/kg BW in mallards (acute oral LD50)

Sensitivity of mammals to paraquat was variable, owing to inherent differences in interspeciesresistance Representative mammals were measurably affected at aerosol concentrations of 0.4 to6.0 µg/L, acute oral doses of 22 to 35 mg/kg BW, dietary concentrations of 85 to 100 mg/kg ration,and drinking water levels of 100 mg/L

22.5.2 Terrestrial Plants and Invertebrates

In terrestrial plants, paraquat’s action is at the point of local absorption (Anonymous 1963).Characteristic damage signs to susceptible species include wilting and general collapse in herbaceousplants Regrowth may occur in some perennial plants, but in resistant species temporary scorchmay be the most marked effect (Anonymous 1963) In sugarcane (Saccharum officinarum), paraquat

application severely desiccated the plant within 72 h, and disrupted activity of leaf amylase andsucrose (Haley 1979) Paraquat, once absorbed in plants, is likely to persist (Bauer 1983) Theaddition of cationic or nonionic surface active agents increases the phytocidal effectiveness ofparaquat (Anonymous 1963), but in combination with various herbicides paraquat was markedlyless phytotoxic to certain cereal grains (O’Donovan and O’Sullivan 1986).

Paraquat adsorbed to soils is usually unavailable to crops In the case of wheat (Triticum aestivum), effects from contaminated soils were negligible until soil residues surpassed 600 to

1000 kg/ha, causing growth reduction of 10%, or 1650 kg/ha, causing elevated residues in leavesbut not in grain (Moyer and Lindwall 1985)

Three species of grains (barley, Hordeum vulgare; wheat; oat, Avena sativa) died (>95% kill)following application of 0.28 kg paraquat/ha (O’Donovan and O’Sullivan 1986) At 0.6 kg/ha,paraquat inhibited germination and growth in seeds of six species of grasses (Kentucky bluegrass,

Poa pratensis; perennial ryegrass, Lolium perenne; bentgrass, Agrostis tenuis; tall fescue, Festuca arundinacea; red fescue, Festuca rubra; orchard grass, Dactylus glomerata), but two species oflegumes (alfalfa, Medicago sativa; red clover, Trifolium pratense) were comparatively resistant(Salazar and Appleby 1982) Paraquat was phytotoxic to several species of terrestrial plants (rape,

Brassica rapa; ryegrass; white clover, Trifolium repens) for several days following application of1.1 to 2.2 kg/ha (Summers 1980) Transpiration rate of soybean (Glycine max) was lowered at

1 mg/kg (Haley 1979) Paraquat is not considered to be carcinogenic or teratogenic, but is weaklymutagenic to some plants (e.g., 4.1% chromosomal aberrations in seeds of wheat at 9.3 mg/kg;Haley 1979) Spray solutions containing 0.6 g paraquat/L applied to crowns of eastern red cedar(Juniperus virginiana) killed up to 90% of small trees and up to 30% of large trees; at 0.3 g/L, up

to 60% of small trees were affected (Engle et al 1988) Seedlings of corn (Zea mays) sprayed with0.2% paraquat ion solution for 6 h had decreased rates of total protein synthesis and some polysomedissociation (Wu et al 1988), suggesting that additional research is needed on mutagenicity ofparaquat in plants

Paraquat resistance has been documented in several genera of weeds For example, resistant strains of barley grass (Hordeum glaucum) were first noted in 1982 in Australia; resistantstrains (based on chromosome counts and resistance to paraquat) were confined to a small number

paraquat-of lucerne fields where paraquat had been used consistently for at least 10 years However, thepotential exists for this biotype to be transferred and established in other areas by the movement

of livestock, machinery, hay, and seeds (Islam and Powles 1988; Tucker and Powles 1988) resistant strains of weeds have been reported in England, Japan, Egypt, and Australia (Polos et al.1988) Paraquat-resistant strains of bacteria, ferns, and other species of flora have been documented(Carroll et al 1988) Paraquat-tolerant ferns (Ceratopteris richardii) were 10 to 20 times moreresistant than sensitive wild-type strains (Carroll et al 1988) Paraquat-resistant strains of perennialrye were up to 10 times more resistant than normal susceptible strains (Faulkner and Harvey 1981)

Paraquat-In the case of barley grass, survival was reduced 50% at 0.025 kg/ha in normal susceptible biotypes;but in resistant biotypes, 3.2 kg/ha were required (Islam and Powles 1988; Tucker and Powles1988) Paraquat-tolerant plants may enjoy certain advantages over nonresistant plants, includingresistance to various air pollutants For example, paraquat-tolerant tobacco plants (Nicotiana tabacum), which had higher superoxide dismutase activity than controls, were tolerant to aerosolsprays of 2 mg SO2/L, while controls experienced severe damage (Tanaka et al 1988)

In every case of resistance, paraquat had been applied 2 or 3 times annually during the preceding

5 to 11 years In some cases, a cross-resistance to atrazine was also reported (Polos et al 1988).Paraquat-resistance mechanisms in plants include increased epicuticular wax (preventing penetra-tion), binding of paraquat to cell walls, restricted movement into chloroplasts, and altered redoxpotential (Polos et al 1988) For example, sequestration of paraquat within the apoplast of the leafseems to be inheritable and controlled by a single nuclear gene with incomplete dominance (Islamand Powles 1988) Studies with paraquat-tolerant strains of various plants, including perennial ryeand tobacco, suggest that tolerance is related to their general ability to rapidly detoxify the generated

oxygen species through increased levels of superoxide dismutase, glutathione reductase, and other

antioxidants (Shaaltiel et al 1988)

At recommended field concentrations, paraquat had negligible effect on soil microflora or soil

fertility, although it did cause a temporary suspension of soil nitrification (Haley 1979) A

concen-tration as low as 1.0 mg/L completely inhibited ammonium and nitrite oxidation for 40 days in a

mixed culture of nitrifying bacteria isolated from soil (Gadkari 1988) Paraquat at 1.6 mg/L

adversely affected Escherichia coli in 6 h, as judged by diminished growth rate and inhibited

synthesis of RNA and protein; at a higher concentration of 18.6 mg/L, interference with metabolism

of glucose and DNA synthesis was observed (Davison and Papirmeister 1971) Four species of soil

bacteria had 50% growth inhibition at paraquat concentrations between 93 and 18,600 mg/kg soil;

moreover, the mode of action in some species of microorganisms may differ from the generally

accepted mechanisms for paraquat toxicity in mammals (Carr et al 1986) Sensitive species of soil

fungi experienced marked growth inhibition between 10 and 25 mg paraquat/kg soil (Summers

1980) In various genera of soil fungi (Rhizopus, Ophiobolus, Helminthosporium, Fusarium,

Euro-tium), paraquat concentrations up to 100 mg/L could be tolerated At higher concentrations, spore

germination was suppressed, mycelial growth was inhibited, and spore development was abnormal

(Haley 1979)

Terrestrial invertebrates show varying degrees of sensitivity to paraquat In honey bees (Apis

mellifera), 100 mg paraquat/kg syrup (diet) produced toxic signs, 4.4 kg/ha applied as a spray killed

90% in 3 days, and 1000 mg/L in drinking water killed most in a few days and 100% within

5 weeks (Summers 1980) In soils, adsorbed paraquat may be ingested by soil invertebrates, such

as earthworms, but it was not absorbed from the gut into tissues and was rapidly lost when the

earthworms were transferred to clean soil (Calderbank 1975) For example, earthworms (Lumbricus

terrestris) fed soil treated with 112 kg paraquat/ha had 111 mg paraquat/kg in gut contents, but

<0.3 mg/kg in the carcass without gut (Summers 1980) Two species of collembolid insects

(Folsomia candida, Tullbergia granulata) fed diets containing 600 mg paraquat/kg for 22 weeks

survived without measurable adverse effects, but higher dietary levels of 1000 and 5000 mg/kg

were associated with decreased survival, lengthier instar development, decreased egg production,

and decreased egg viability (Subagja and Snider 1981) Adults and larvae of the German cockroach

(Blattella germanica) died after consuming diets containing 1000 mg paraquat/kg (Summers 1980)

Also, paraquat was lethal to two species of mites (Tetranchus urticae, Typhlodromus sp.) at

concentrations below recommended field application rates (Summers 1980)

22.5.3 Aquatic Organisms

In general, paraquat is more toxic to aquatic fauna in soft water than in hard water, more toxic

to early developmental stages than to juveniles or adults, and more toxic in formulations containing

wetting agents than in formulations without these agents (Summers 1980; Arunlertarce and Kawatsu

1992) In water, paraquat is taken up rapidly by plants or adsorbed to particulate matter in the water

column; however, paraquat is not bioconcentrated by aquatic fauna (Calderbank 1975; Summers

1980) Paraquat effects on aquatic biota are summarized in Table 22.3, and these data suggest

several trends Early developmental stages of certain species of crustaceans are extremely sensitive,

and significant adverse effects occur in the range of 0.9 to 100 µg/L, although most species of

crustaceans and all other species of invertebrates tested were relatively unaffected at concentrations

below 1000 µg/L Freshwater algae and macrophytes are eliminated after treatment with 250 to

500 µg/L, but marine algae are relatively resistant and require 5000 µg/L or higher to produce

significant growth inhibition Aquatic vertebrates usually are not adversely affected and show little

accumulation at 1000 µg/L or lower; but at 500 µg/L, frog tadpoles have low survival and a high

frequency of developmental abnormalities, and carp experience biochemical upset

Paraquat controlled Typha and Phragmites weeds in Egyptian irrigation canals, drains, and

marshes without apparent harm to fishes (Haley 1979) Paraquat residues in decomposed plants

become available for adsorption to sediments and bottom muds and are not readily available for

microbial degradation (Summers 1980) Indirect fish kills may occur from anoxia due, in part, to

consumption of dissolved oxygen by decaying weeds (Bauer 1983) Paradoxically, it has been

suggested that paraquat may be helpful in improving the oxygen status of aquatic environments at

a concentration of 1 mg/L by restricting nitrate production due to inhibition of bacterial nitrification

(Chan and Leung 1986; Gadkari 1988) At effective herbicidal concentrations, paraquat was also

toxic to eggs, but not adults, of three species of gastropod vectors of bilharzia (Bulinus truncatas,

Biomphalaria alexandrina, Lymnaea calliaudi); newly hatched snails were the most sensitive (Haley

1979)

Changes in fauna of a reservoir following use of paraquat for weed control are likely to be

indirect effects caused by decomposition of angiosperms (Brooker and Edwards 1974) Planktonic

invertebrates closely associated with aquatic macrophytes were either eliminated by paraquat or

survived at lower densities for at least a year posttreatment; analysis of fish stomachs showed

dietary changes following weed control and reflected availability of many invertebrate species

associated with aquatic plants (Brooker and Edwards 1974)

Paraquat can induce activities of antioxidant enzymes such as superoxide dismutase, glutathione

peroxidase, and catalase in many species of plants, invertebrates, and vertebrates Results of studies

with ribbed mussels (Geukensia demissa) support the hypothesis that these bivalve molluscs can

activate redox cycling compounds and demonstrate responses typical of oxidative stress observed

in other species (Wenning et al 1988) Paraquat also disrupts glucose metabolism and

acetylcho-linesterase activity and accumulates in melanin Disrupted glucose metabolism in paraquat-stressed

carp was attributed to a high level of circulating epinephrine (Simon et al 1983) Paraquat-induced

acetylcholinesterase inhibition in erythrocytes and electric organs of the electric eel (Electrophorus

electricus) was reversible (Seto and Shinohara 1987,1988) Paraquat tended to concentrate in

melanin, as judged by accumulation in neuromelanin of frogs (Rana temporaria) after

intraperito-neal injection (Lindquist et al 1988), with important implications for research on Parkinson’s

disease It seems that paraquat has a structural similarity to a metabolite of

1-methyl-4-phenyl-1,2,3, 6-tetrahydropyridine (MPTP), which may induce a Parkinson-like condition MPTP and its

metabolites, like paraquat, have melanin affinity (Lindquist et al 1988)

Table 22.3 Effects of Paraquat on Selected Species of Aquatic Plants and Animals (Concentrations

are in mg of paraquat cation per liter of medium.)

Taxonomic Group, Organism,

and Other Variables

Concentration (mg/L) and Effect Reference a ALGAE AND MACROPHYTES

Eurasian watermilfoil, Myriophyllum

spicatum

1.0; residues up to 112 mg/kg DW in 14 days 4, 5

Cattail, Typha latifolia 0.5; shoreline colonies severely affected after

exposure for 32 days

3

Duckweed, Spirodella oligorrhiza 0.5; inhibits chlorosis in 48 h 3 Marine algae

INVERTEBRATES

Mud crab

Isopod, Asellus meridianus

Freshwater invertebrates, 3 species

(Asellus, Lymnaea, Sialis)

0.5; no deaths in 4 days following spray application

to British lake

3 Gastropod

parts, after exposure for 3 days ranged between 1.5 and 2.8, and were dose dependent

8

Hermit crab

exposure for 3 days ranged between 3.2 and 15, and were dose dependent

8

Brown shrimp, Penaeus aztecus >1.0; 50% immobilization in 48 h 6

American oyster, Crassostrea virginica >1.0; 50% growth reduction in 96 h 6

Louisiana red crayfish, Procambarus

clarkii

All stages Sublethal; Dose-dependent increase in

hyperactivity and oxygen consumption

9 Daphnid

Cladoceran

Liver fluke, Fasciola hepatica

Egg through miracidium 5; LC50 (20 days); effects counteracted by dinoseb 12

of miracidia

12

Table 22.3 (continued) Effects of Paraquat on Selected Species of Aquatic Plants and Animals

(Concentrations are in mg of paraquat cation per liter of medium.)

Taxonomic Group, Organism,

and Other Variables

Concentration (mg/L) and Effect Reference a

transferred to paraquat-free medium died within

48 h

14

transferred to paraquat-free medium died within

24 h

14

Ribbed mussel,

Geukensia demissa 93; elevated catalase activity, lipid peroxidation rate,

and total superoxide dismutase levels in 12–36 h

FISHES

Common carp

Cyprinus carpio 0.5; after 6 days, 300% increase in phosphorylase

and 200% increase in glucose-6-phosphatase activities in liver; increase in sugar level and serum lactic dehydrogenase activity

18

and in blood sugar levels during exposure for

6 days; effects enhanced by the herbicide methidation

19

activity which gradually increased to 130% of control values during exposure for 2 weeks, suggesting that paraquat may influence resynthesis of acetylcholinesterase

28

C carpio 10; during exposure for 96 h, significant alterations

were recorded in lipid peroxidation rate, hemoglobin concentration, and erythrocyte antioxidant enzymes, that is, catalase, superoxide dismutase, and glutathione peroxidase activities

20

gill, liver, and brain; effects exacerbated under conditions of hypoxia

32

embryo developmental stages, respectively

31

after exposure for 2 h in serum (13 mg/L), heart (39 mg/L), muscle (102 mg/L), and brain (214 mg/L)

21

Table 22.3 (continued) Effects of Paraquat on Selected Species of Aquatic Plants and Animals

(Concentrations are in mg of paraquat cation per liter of medium.)

Taxonomic Group, Organism,

and Other Variables

Concentration (mg/L) and Effect Reference a

C carpio 67.5–134.1; LC50 (96 h) values for fry and

fingerlings, respectively

31 Plecostomid catfish

Plecostomus commersonii, fry 0.6; decrease in cardiac contraction rate after

4 species 1.0; maximum residues, in mg/kg whole-body FW,

during exposure for 16 days ranged between 0.6 in

green sunfish (Lepomis cyanellus) and 1.6 in bluegill (Lepomis macrochirus); intermediate

values were recorded for rainbow trout

(Oncorhynchus mykiss) and channel catfish (Ictalurus punctatus)

22

3 species 10; in exposures lasting up to 96 h, paraquat

caused an exposure-dependent increase in lipid peroxidation rate and in activity enhancement of peroxide metabolism enzymes in erythrocytes

23

3 species 250; lesions in skin and gills of survivors and

increasing susceptibility to secondary invaders

30

Smallmouth bass, Micropterus dolomieui 1.0; adverse sublethal effects 22 Striped mullet

Mugil cephalus 1.0; LC50 (16 days); survivors had pronounced gill

histopathology and residues, in mg/kg FW, of 0.2 in muscle, 0.2 in ovary, 4.7 in skin, and 6.1 in digestive tract

P gonionotus 4.0; gill histopathology evident after exposure for

12 days; gills normal during exposure for only

Poeciliid fish, Cnesterodon

decemmaculatus; fry

Medaka, Oryzias latipes

Rainbow trout

Channel catfish, Ictalurus punctatus >100; LC50 (96 h) 5, 10, 11

Table 22.3 (continued) Effects of Paraquat on Selected Species of Aquatic Plants and Animals

(Concentrations are in mg of paraquat cation per liter of medium.)

Taxonomic Group, Organism,

and Other Variables

Concentration (mg/L) and Effect Reference a