Toxicity of pesticides to aquatic microorganisms: A review

Authors

  • Marie E. DeLorenzo,

    Corresponding author
    1. U.S. Department of Commerce/NOAA, National Ocean Service, Center for Coastal Environmental Health and Biomolecular Research, 219 Fort Johnson Road, Charleston, South Carolina 29412
    • U.S. Department of Commerce/NOAA, National Ocean Service, Center for Coastal Environmental Health and Biomolecular Research, 219 Fort Johnson Road, Charleston, South Carolina 29412
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  • Geoffrey I. Scott,

    1. U.S. Department of Commerce/NOAA, National Ocean Service, Center for Coastal Environmental Health and Biomolecular Research, 219 Fort Johnson Road, Charleston, South Carolina 29412
    Search for more papers by this author
  • Philippe E. Ross

    1. Colorado School of Mines, Environmental Science and Engineering Division, Golden, Colorado 80401, USA
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Abstract

Microorganisms contribute significantly to primary production, nutrient cycling, and decomposition in estuarine ecosystems; therefore, detrimental effects of pesticides on microbial species may have subsequent impacts on higher trophic levels. Pesticides may affect estuarine microorganisms via spills, runoff, and drift. Both the structure and the function of microbial communities may be impaired by pesticide toxicity. Pesticides may also be metabolized or bioaccumulated by microorganisms. Mechanisms of toxicity vary, depending on the type of pesticide and the microbial species exposed. Herbicides are generally most toxic to phototrophic microorganisms, exhibiting toxicity by disrupting photosynthesis. Atrazine is the most widely used and most extensively studied herbicide. Toxic effects of organophosphate and organochlorine insecticides on microbial species have also been demonstrated, although their mechanisms of toxicity in such nontarget species remain unclear. There is a great deal of variability in the toxicity of even a single pesticide among microbial species. When attempting to predict the toxicity of pesticides in estuarine ecosystems, effects of pesticide mixtures and interactions with nutrients should be considered. The toxicity of pesticides to aquatic microorganisms, especially bacteria and protozoa, is an area of research requiring further study.

INTRODUCTION

Microorganisms are important inhabitants of aquatic ecosystems, where they fulfill critical roles in primary productivity, nutrient cycling, and decomposition. Aquatic environments receive direct and indirect pesticide inputs, inevitably exposing microorganisms to pesticides. While pesticides elicit a variety of acute and chronic toxicity effects in microorganisms, microorganisms also have the capability to accumulate, detoxify, or metabolize pesticides to some extent. Detrimental effects of pesticides on microbial species may have subsequent impacts to higher trophic levels. For example, changes in the macromolecular composition of phytoplankton species or shifts in community composition can affect the growth rate of zooplankton grazers [1].

Estuaries serve as critical feeding and nursery grounds for many aquatic organisms, including commercially and recreationally important fish and shellfish species. These productive, diverse ecosystems are particularly vulnerable to pollution because they serve as repositories for pollutants from upland sources. Millions of pounds of active pesticide ingredients are applied in coastal watersheds each year. Pesticide usage in the South Florida Water Management District, which is adjacent to the Everglades National Park and Florida Bay (FL, USA), includes an estimated 1,415 tons of atrazine, 36 tons of en-dosulfan, and 622 tons of chlorpyrifos per year [2]. Runoff of agricultural pesticides into estuaries poses significant toxico-logical risks to resident organisms [3].

Toxicity data involving microorganisms and pesticides are limited. Most studies have focused on microbial degradation of pesticides rather than impacts on natural microbial populations. In addition, studies of pesticide effects on soil microbes are far more common than studies of those in aquatic environments. Data involving marine or estuarine microorganisms are even scarcer. This paper will provide a review of studies describing pesticide toxicity to aquatic microorganisms. It will describe the biochemical mode of action of some pesticides, compare the sensitivities of microbial species to various pesticides, comment on the ecological relevance of the findings, and define areas where more research is needed.

MECHANISMS OF PESTICIDE ACTION

Pesticides can be classified according to their mechanisms of action. For example, organochlorine, organophosphate, and carbamate insecticides act primarily by disrupting nervous system function, while herbicides target mainly photosynthesis pathways (Table 1). The mechanism of pesticide action in microbial species may not be the same as for the target organisms. In microorganisms, pesticides have been shown to interfere with respiration, photosynthesis, and biosynthetic reactions as well as cell growth, division, and molecular composition.

Over half the herbicides in current use act primarily on the light reaction of photosynthesis. Many groups of herbicides act by inhibiting the Hill reaction of electron transport, including ureas, cyclic ureas, triazines, acylanilides, phenylcar-bamates, and triazinones. The bipyridinium herbicides, such as diquat and paraquat, act by intercepting electrons from the reducing side of photosystem I. The herbicidal cation is reduced by the light reactions of photosynthesis to form a relatively stable free radical. In the presence of oxygen, the bi-pyridinium free radical becomes oxidized to form the original ion, which is then free to react again, and an activated oxygen species is formed that destroys the cell tissue. Bipyridinium herbicides are effective only in the presence of light [4].

Fluometuron is a herbicide with two independent mechanisms of action. As a substituted phenylurea compound, it inhibits photosynthesis. In addition, it inhibits carotene biosynthesis, a process known as bleaching. Chlorophyll formation is not inhibited directly, but the pigment is destroyed in the presence of light because of the missing photo-oxidative carotene shield. In the unicellular algae Chlorella pyrenoidosa and Euglena gracilis, this was demonstrated by normal chlorophyll formation in the presence of fluometuron in the dark but not in the light [5]. Pyridazinones are other compounds that interfere with carotenoid accumulation. They also interfere with chloroplast lipids by altering their fatty acid composition [4]. Chloroacetamide herbicides, such as alachlor and meta-zachlor, are thought to act by disrupting the synthesis of fatty acids, resulting in the inhibition of cell division and growth [6].

The nitrodiphenyl ether herbicide 5-[2-chloro-4-(trifluoro-methyl)phenoxy]-2-nitroacetophenone oxime-O-(acetic acid, methyl ester) (DPEI) induced light- and oxygen-dependent lip-id peroxidation and chlorophyll bleaching in the green alga Scenedesmus obliquus. A study of the mode of action of ni-trodiphenyl ether herbicides by Bower et al. [7] revealed that under conditions of low oxygen, the toxic effects of DPEI were reduced by prometyne and 3-(3,4-dinitrophenyl)-1,1-di-methylurea (DCMU), both of which are inhibitors of photo-synthetic electron transport. Under high oxygen conditions, DCMU did not protect S. obliquus cells from chlorophyll bleaching induced by DPEI but did protect against paraquat. The DPEI, but not paraquat, induced tetrapyrole formation in treated cells in the dark. This was also observed in a mutant lacking photosystem I but was suppressed under low oxygen conditions. Bower et al. [7] concluded that during diphenyl ether toxicity in S. obliquus, photosynthetic electron transport functions to maintain high oxygen concentrations through water splitting in the algal suspension. With paraquat, photosyn-thetic electron transport reduces the herbicide to a radical species, which initiates lipid peroxidation. Another method of herbicide action is interference with microtubular systems. The microtubule-forming unit is the dimeric protein tubulin. The pesticide colchicine binds to the tubulin dimer and inhibits microtubule formation. Loss of spindle microtubules affects nuclear division and the separation of chromosomes. Lack of cortical microtubules results in disrupted morphogenesis of cells and tissues. Other antimicrotubular herbicides include dinitroanilines, carbamates, phosphoric amides, chlorthaldi-methyl, propyzamide, and terbutol [8].

Amiprophos-methyl, like dinitroanilines, affects flagellar regeneration. Simultaneously, the synthesis of new tubulin and one additional flagellar protein is inhibited. Amiprophos-meth-yl disrupts control of cellular calcium ion concentrations, possibly causing the inhibition of tubulin synthesis [9].

The p-nitrophenol is primarily used in the manufacture of the organophosphate insecticide parathion. The p-aminophenol is a degradation product of p-nitrophenol. These nitrosubsti-tuents of phenol, but not phenol itself, were found to inhibit the production of chlorophyll a, protein, and carbohydrate in the cyanobacterium Nostoc linckia. The 14CO2 uptake and activities of nitrate reductase, nitrogenase, and glutamine synthetase were also hampered by these nitrophenols. Photohet-erotrophic conditions (0.1% acetate) did not reduce the toxicity of the nitrophenols. The addition of adenosine triphosphate, however, did reverse the toxicity [10]. In another study, p-aminophenol concentrations above 2 mg/ml were found to inhibit cell number, chlorophyll a, total carbohydrate production, 14CO2 uptake, and nitrate reductase and nitrogenase activity in Chlorella vulgaris, N. linckia, and Nostoc muscorum. The adverse effects of the parent compound, p-nitrophenol, were found to be alleviated in the presence of p-aminophenol at concentrations <2 mg/L [11]. In both studies, transmission electron microscopy revealed many cytological abnormalities, such as secretion of mucus around the filament and induction of spore formation in the algae subjected to selected phenols, indicating that the toxicants directly interfere with membrane properties and enzymes.

Chlorophenols are often used as broad-spectrum biocides. They are potent uncouplers of oxidative photophosphorylation [12]. Pentachlorophenol has several sites of action, including photophosphorylation, protein synthesis, and lipid biosynthesis [13].

The mechanism by which tributyltin affects algae as well as higher organisms is the inhibition of respiratory function [14]. The trialkyl tins are thought to act on mitochondria by inhibiting the adenosine triphosphase responsible for adenosine triphosphate synthesis [4].

A study of the toxicity of DDT to algae revealed that p,p′-DDT stimulated photorespiration and suppressed the incorporation of 14C from 14CO2 into the C4-dicarboxylic acid pathway. The DDT-induced shift in metabolism from an efficient to a nonefficient pathway was most likely through disruption of cyclic photophosphorylation [15]. For many other pesticides, although deleterious effects have been documented in microorganisms, the mechanism of toxicity to such nontarget organisms remains unknown.

Table Table 1.. Summary of pesticide mechanisms of action on target organisms
Pesticide classGroups includedGeneral toxic effectSpecific site of action
OrganophosphatesCarbamatesNervous system inhibitionAcetylcholinesterase
OrganochlorinesCyclodienesNervous system inhibitionGABA receptor
HerbicidesUreas, cyclic ureas, triazines, acylanilides, phenylcarbamates, triazinonesPhotosynthesis inhibitionHill reaction of electron transport
 BipyridiniumsPhotosynthesis inhibition (light reaction)Reducing side of photosystem I
 PyridazinonesBiosynthesis inhibitionCarotene accumulation
 ChloroacetamideBiosynthesis inhibitionFatty acid synthesis
 Dinitroanilines, phosphoric amides, chlorthaldimethyl, propy-zamide, cholchicine, terbutolBiosynthesis inhibitionMicrotubule formation
Broad-spectrum biocidesChlorophenolsMultiple inhibiting actionsPhophosphorylation, protein synthesis, lipid biosynthesis
 Tributyl tins, trialkyl tinsRespiratory system inhibitionMitochonrial ATPase

MICROBIAL SENSITIVITY TO PESTICIDES

It is not surprising that there is considerable diversity in the sensitivity of microorganisms to pesticides. Microorganisms span three kingdoms, including > 50,000 different species of bacteria, algae, and protozoa. Microorganisms cover a tremendous range of size classes and morphologies, exist in every imaginable habitat, and include multiple feeding types, mobile, and nonmobile forms, and a wide variety of reproductive strategies and growth rates. The majority of the available pesticide data regarding aquatic microorganisms is for algae. Of that data set, most of the pesticides studied are herbicides, and of those herbicides, atrazine is the most extensively tested compound (Table 2). Far fewer pesticide studies exist for aquatic bacteria and protozoa. In the algal toxicity tests described in the following, most EC50s were determined using growth rate as an end point. Generally, algal growth rate was assessed using direct cell counts or optical density. The duration of the tests varied, with 24 to 96 h being the most common exposures. Studies of microorganisms from all aquatic ecosystems were included with attention drawn to those involving marine and estuarine species (Table 2). The studies are grouped by class of pesticide and include data for pesticides that are no longer registered for use in the United States yet are still routinely detected in surface waters (e.g., DDT).

Chlorinated hydrocarbons

Various responses were observed among marine and freshwater algae to the effects of chlorinated hydrocarbons. Aldrin, dieldrin, and endrin had no significant effect on respiration of green and bluegreen algae at concentrations of up to 1 mg/L [16]. At higher concentrations (100 mg/L), aldrin and dieldrin lowered adenosine triphosphate levels but not population densities of algae [17]. In general, aldrin is the least inhibitory of the three compounds, with marine algae usually having greater tolerance than freshwater forms [18]. Lee et al. [15] found that concentrations of DDT below 10 μg/L were found to inhibit photosynthesis in marine planktonic algae. At concentrations between 3.6 and 36 μg/L p,p′-DDT also inhibited photosyn-thetic CO2 fixation in the green alga Selenastrum capricor-nutum. Chlorella sp. was affected by less than 0.3 μg/L DDT. The bluegreen alga Anabaena sp. was found to be sensitive to DDT at 1 mg/L [19]. Another organochlorine, mirex, had no effect on marine algae at 0.2 μg/L, although it was concentrated up to 10,000 times by the algae [20]. In another study, 100 μg/L of both mirex and methoxychlor only slightly inhibited algal growth (19 and 17% less than control, respectively). A mixture of mirex and methoxychlor (each at 50 μg/L) had no effect on population growth [21].

In a study of the comparative effects of chlorophenols, toxicity values (96-h EC50) in S. capricornutum were found to decrease from 150 mg/L for phenol to 0.42 mg/L for pen-tachlorophenol. Thus, toxicity increased with increasing number of substituted chlorine atoms [12].

Endosulfan

Endosulfan is a synthetic, chlorinated cyclodiene insecticide (same chemical group as DDT). It is the only remaining pesticide of this class still used in the United States. It is applied to grains, tea, fruits, vegetables, tobacco, and cotton.

Endosulfan had detrimental effects on reproduction in the green alga Chlamydomonas reinhardtii. A single exposure during the four developmental stages of the sexual life cycle delayed meiosis for 5 d. Gamete production and zygote formation were unaffected. An endosulfan concentration of 10.17 mg/L, applied to pregametogenesis cells, lessened meiotic division 35% relative to controls. The number of first-day dividing zygotes was reduced 100% after treatment with endosulfan [22]. In the marine red alga Champia parvula, endosulfan concentrations of 47 and 130 μg/L chronically impaired female growth and tetrasporophytes, respectively. Higher levels, 360 to 600 μg/L, produced chronic reproductive effects in the alga. The maximum allowable toxicant concentration for endosulfan in C. parvula was <47 μg/L [23].

Endosulfan at 10 mg/L reduced C. vulgaris growth rates by 15%, and growth was completely inhibited in this alga at 100 mg/L endosulfan. A 10-d LC50 of 41.5 mg/L was determined. In another alga, Anabaena doliolum, growth was reduced at all endosulfan concentrations tested (0.1–100 mg/L), and growth stopped completely at concentrations greater than 3 mg/L. The 10-d LC50 for A. doliolum was 2.15 mg/L [24].

Endosulfan inhibited growth in the bluegreen algae Anabaena sp. and Aulosira fertilissima. Anabaena sp. cultures were reduced by 51.7% and A. fertilissima by 37.7% at 1 mg/L endosulfan [19]. Photosynthesis in A. fertilissima was inhibited 40.2% at 20 mg/L endosulfan [25]. Rao and Lal [26] found that Anabaena sp. and A. fertilissima rapidly accumulated and concentrated endosulfan. The researchers also found that endosulfan was biotransformed by these bluegreen algae to endosulfan ether and endosulfan lactone. Growth of the bluegreen alga Westiellopsis prolifica was inhibited by 10 mg/L endosulfan, and nitrogen fixation was reduced 93% at 100 mg/L endosulfan [27]. Mohapatra and Mohanty [24] found endosulfan to be more toxic to the cyanobacterium Anabaena sp. than to the green alga Chlorella sp. A delayed inhibition of growth exhibited by Chlorella sp. was thought to be due to efficient accumulation and concentration of endosulfan inside the cell. Both algae were more sensitive to endosulfan than to the organophosphate pesticide dimethoate.

Rajendran et al. [28] examined the effects of endosulfan on heterotrophic bacterial populations in water and sediment samples of the Vellar estuary, South India. The percentage of the water bacterial population inhibited by endosulfan ranged from 30.99% at 0.002 mg/L to 70.58% at 2 mg/L. The majority of the endosulfan resistant bacteria were Pseudomonas spp. Bacteria isolated from water samples were more sensitive to endosulfan than those isolated from sediment.

In a study of estuarine microbial communities, DeLorenzo et al. [29] found that bacterial abundance was significantly reduced at 1 and 10 μg/L endosulfan, while heterotrophic bacterial productivity was not. Phototrophic biovolume was also reduced at 1 and 10 μg/L endosulfan, and the abundance of several cyanobacterial species declined. There was no significant effect on heterotrophic ciliate or flagellate abundance.

Organophosphate insecticides

Christie [30] observed that 100 mg/L malathion had little significant effect on the green alga Chlorella pyrenoidosa, and cultures of bluegreen and green algae were not adversely affected by accumulated levels (50–72 mg/L) of parathion. Parathion was found, however, to cause reproductive inhibition at 7.86 g/L in Chlorella fusca [31]. The nitrogen-fixing bluegreen algae Cylindrospermum sp. and Aulosira fertilissima grew well in the presence of 300 and 400 mg/L diazinon, respectively [32].

Table Table 2.. Summary list of pesticide toxicity tests involving aquatic microbial species. Not all atrazine studies are included (see Huber [65] and Solomon et al. [66] for reviews). OP = organophosphate. OC = organochlorine, TC = thiocarbamate, BP = bipyridinium
PesticidePesticide classSpecies nameType of organismTest end pointReference
2,4-DPhenoxy herbicideSeveral algal speciesFilamentous7-d chlorophyll production[33]
  Chlorella fuscaGreen algaReproduction[31]
  Vibrio fisheriMarine bacterium5-min EC50, light reduction[39]
2,4,5-TPhenoxy herbicideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
AcroleinCarboyl pesticideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  Aphanizomenon flosaquaeCyanobacterium7-d carbon uptake[51]
  Scenedesmus quadricaudaGreen alga7-d carbon uptake[51]
  Selenastrum capricornutumGreen alga7-d carbon uptake[51]
AlachlorChloroacetamide herbicideAlgal communityFreshwaterChlorophyll a, composition, densities Algal biovolume[60]
  Benthic algal communityFreshwater [61]
  Pavlova sp.PyrmnesiophyteGrowth rate[62]
AnilazineTriazine herbicideChlorella fuscaGreen algaReproduction[31]
AtrazineTriazine herbicideChlorella vulgarisGreen alga7-d chlorophyll[33]
  Stigeoclonium tenueGreen alga7-d chlorophyll[33]
  Oscillatoria luteaCyanobacterium7-d chlorophyll[33]
  Scenedesmus sp.Green algaGrowth rate[67]
  Anabaena flos-aquaeCyanobacterium3, 5, 7-d growth rates[69]
  Skeletonema costatumDiatom5-d growth, fluorescence[70]
  Skeletonema costatumMarine diatom48-h growth rate[40]
  Minutocellus polyorphusMarine diatom48-h growth rate[40]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
  Nitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon intake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  C. reinhardtiiGreen algaGrowth rate[75]
  S. quadricaudaGreen algaGrowth rate[75]
  Scenedesmus subspicatusGreen alga96-h growth rate[73]
  Pavlova sp.Prymnesiophyte96-h growth rate[62]
  Phaeodactylum tricornutumDiatomGrowth rate[68]
  Nannochloris oculataCyanobacteriumGrowth rate[68]
  Chlamydomonas sp.Green alga1-h EC50 02 production[76]
  Cyclotella nanaMarine diatom1-h EC50 02 production[76]
  Navicula insertaMarine diatom1-h EC50 02 production[76]
  Thalassiosira fluviatilisEstuarine diatom7-d photosynthesis, cell density[77]
  Nitzschia sigmaEstuarine diatom7-d photosynthesis, cell density[77]
  Cryptomonas sp.CryptophytePopulation density[78]
  Mallomonas sp.CryptophytePopulation density[78]
  Benthic algal communityFreshwaterAlgal biovolume[61]
  Microbial communitiesEstuarine72-h composition, density, C uptake, chlorophyll a[29]
  Chlamydomonas sp.Green alga28-d growth rate[74]
  Chlorella sp.Green alga28-d growth rate[74]
  Pediastrum sp.Green alga28-d growth rate[74]
  Scenedesmus sp.Green alga28-d growth rate[74]
  Cyclotella gammaDiatom28-d growth rate[74]
  Cyclotella meneghinianaDiatom28-d growth rate[74]
  Synedra acusDiatom28-d growth rate[74]
  Synedra radiansDiatom28-d growth rate[74]
  Pond communityFreshwater microcosmO2 production/dissolved nitrate[85]
AzocyclotinOrganotinMicrobial communityFreshwater microcosmPhytoplankton densities[53]
BromacilPhenylurea herbicideC. reinhardtiiGreen algaResistance[8]
CarbarylOP insecticideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
  Chlorella pyrenoidosaGreen algaPopulation density[30]
  Nostoc muscorumCyanobacteriumGrowth rate[19]
  CylindrospermumCyanobacteriumGrowth rate[19]
  Nitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
CarbofuranCarbamate insecticideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
ChlorpyrifosOP insecticideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
  Anabaena sp.CyanobacteriumGrowth rate[19]
  Microbial communitiesEstuarine72-h composition, density, C uptake, chlorophyll a[29]
  Skeletonema costatumMarine diatom48-h growth rate[40]
  Minutocellus polyorphusMarine diatom48-h growth rate[40]
  Pond communityFreshwater algaePopulation density[48]
ChlortoluronPhenylurea herbicideChlorella fuscaGreen algaReproduction[31]
CyanazineTriazine herbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
DDEOC insecticideThalassiosira pseudonanaDiatomGrowth rate[109]
p.p′-DUIOC insecticideS. capicornutumGreen algaPhotosynthetic CO2 fixation[19]
  Chlorella sp.Green algaPhotosynthetic CO2 fixation[19]
  Anabaena sp.CyanobacteriumPhotosynthetic CO2 fixation[19]
DiazinonOP insecticideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
  Cylindrospermum sp.CyanobacteriumGrowth rate[32]
  Aulosira fertilissimaCyanobacteriumGrowth rate[32]
DimethoateOP insecticideChlorella vulgarisGreen algaGrowth rate[24]
  Anabaena doliolumCyanobacteriumGrowth rate[24]
  S. incrassatulusGreen algaGrowth, density, macromolecules, pigments[36]
DiquatBP herbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillaoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
  Microbial communitiesFreshwaterCell density, biomass, community composition[56]
DiuronPhenylurea herbicideC. reinhardtiiGreen algaResistance[8]
  Bumilleriopsis filiformisGreen algaResistance[8]
  Euglena gracilisGreen algaResistance[8]
EndosulfanCyclodiene insecticideC. reinhardtiiGreen algaMeiotic division[22]
  Champia parvulaRed algaReproduction[23]
  Chlorella vulgarisGreen algaGrowth rate[24]
  Anabaena doliolumCyanobacteriumGrowth rate[24]
  Anabaena sp.CyanobacteriumGrowth rate[19]
  Aulosira fertilissimaCyanobacteriumGrowth rate[19]
  Aulosira fertilissimaCyanobacteriumPhotosynthesis[25]
  Westiellopsis prolificaCyanobacteriumGrowth rate/N fixation[27]
  Bacterial isolatesEstuarinePopulation density[28]
  Microbial communitiesEstuarine72-h composition, density, C uptake, chlorophyll a[29]
FenitrothionOP insecticideChlamydomonas segnisGreen algaGrowth rate/biomass[34]
  Chlorella pyrenoidosaGreen algaGrowth rate/biomass[34]
  Sceneesmus obliquusGreen algaGrowth rate/biomass[34]
  Ankistrodesmus falcatusGreen algaGrowth rate/biomass[34]
  S. capricornutumGreen algaGrowth rate/biomass[34]
  Anabaena sp.CyanobacteriumGrowth rate/biomass[34]
FonofosOP insecticideVirio fisheriMarine bacterium5-min EC50, light reduction[39]
GardoprimTriazine herbicideNatural communityFreshwater algaeGrowth rate[59]
GesapaxTriazine herbicideNatural communityFreshwater algaeGrowth rate[59]
GlyphosateHerbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  F'seudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
  Periphyton communityFreshwater4-h photosynthesis EC50[57]
HexazinoneTriazine herbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
 A. flos-aquaeCyanobacterium7-d carbon uptake [51]
 S. quadricaudaGreen alga7-d carbon uptake [51]
 S. capricorniitumGreen alga7-d carbon uptake [51]
IsofenphosOP insecticideVibrio fisheriMarine bacterium5-min EC50, light reduction[39]
IsoproturonUrea herbicideScenedesmus subspicatusGreen alga96-h growth rate[73]
MalathionOP insecticideChlorella pyrenoidosaGreen algaGrowth rate[31]
  Chlorogloea fritschiiCyanobacteriumGrowth rate[20]
  Stigeoclonium sp.Green algaChlorophyll production[34]
  Tribonema sp.ChrysophyteChlorophyll production[34]
  Vaucheria sp.ChrysophyteChlorophyll production[34]
ManebTC fungicideEuglena gracilisGreen algaGrowth rate[94]
MirexOC insecticideChlorella pyrenoidosaGreen algaGrowth rate[21]
  Chlorococcum sp.Marine green alga7-d growth rate[20]
  Dunaliella tertiolectaMarine green alga7-d growth rate[20]
  Chlamydomonas sp.Marine green alga7-d growth rate[20]
  Nitzschia sp.Marine diatom7-d growth rate[20]
  T. pseudonanaMarine diatom7-d growth rate[20]
  Porphyridium omentumMarine red alga7-d growth rate[20]
MecopropAryloxyalkanoic acid herbicide OC insecticideScenedesmus subspicatusGreen alga96-h growth rate[73]
Methoxyclor Chlorella pyrenoidosaGreen algaGrowth rate[21]
MetribuzinTriazine herbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
MonuronPhenylurea herbicideChlorella sp.Green algaGrowth rate[58]
  C. reinhardtiiGreen algaResistance[8]
NabamTC fungicideEuglena gracilisGreen algaGrowth rate[94]
ParaquatBP herbicidePhormidium sp.CyanobacteriumGrowth rate[56]
ParathionOP insecticideChlorella pyrenoidosaGreen algaGrowth rate[30]
  Vibrio fisheriMarine bacterium5-min EC50, light reduction[39]
  Chlorella fuscaGreen algaReproduction[31]
PentachlorophenolChlorophenolS. capricornutumGreen alga96-h EC50, growth rate[12]
SimazineTriazine herbicideMultiple speciesFreshwaterChlorophyll production[33]
  Chlorella fuscaGreen algaReproduction[31]
  Nitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
TebuthiuronSubstituted urea herbicideNitzschia sp.Diatom7-d carbon uptake[51]
  Cyclotella meneghianaDiatom7-d carbon uptake[51]
  Microcystis aeruginosaCyanobacterium7-d carbon uptake[51]
  Oscillatoria sp.Cyanobacterium7-d carbon uptake[51]
  Pseudoanabaena sp.Cyanobacterium7-d carbon uptake[51]
  Anabaena inaequalisCyanobacterium7-d carbon uptake[51]
  A. flos-aquaeCyanobacterium7-d carbon uptake[51]
  S. quadricaudaGreen alga7-d carbon uptake[51]
  S. capricornutumGreen alga7-d carbon uptake[51]
TemephosOP insecticideChlamydomonas sp.Green algaGrowth rate[37]
  Navicula pelliculosaDiatom7-d growth rate[38]
  Chlorella pyrenoidosaGreen alga7-d growth rate[38]
  Navicula minimaDiatom7-d growth rate[38]
  Coccochloris peniocystisCyanobacterium7-d growth rate[38]
  Oscillatoria sp.Cyanobacterium7-d growth rate[38]
  Chlorella vularisGreen alga7-d growth rate[38]
  Vibrio natrigensBacterium7-d growth rate[38]
ThimetonInsecticideS. incrassatulusGreen algaGrowth, density, macromolecules, pigments[36]
ZinebTC fungicideEuglena gracilisGreen algaGrowth rate[94]

Malathion had a partial inhibitory effect on growth of the bluegreen alga Chlorogloea fritschii, and growth was permanently suppressed at 200 mg/L [19]. Another study showed that malathion inhibited chlorophyll production in Stigeoclon-ium, Tribonema, and Vaucheria by 100% at 1μg/L [33]. This study also provided direct evidence that algae can degrade malathion in the presence of light.

Axenic batch cultures of 12 freshwater algae were used to study the effects of the organophosphorus insecticide fenitro-thion on the freshwater phytoplankton Chlamydomonas seg-nis, Chlamydomonas reinhardtii, Chlorella pyrenoidosa, Chlorella vulgaris, Cosmarium sp., Pediastrum sp., Scene-desmus obliquus, Staurastrum sp., Ankistrodesmus falcatus, Navicula sp., Anabaena sp., and Selenastrum capricornutum. Levels of 10 mg/L significantly reduced growth rate and standing crop in all species. The suggested mode of action was that fenitrothion prevented normal mitotic divisional processes from occurring, resulting in an accumulation of macromolecules and subsequent cell weight increase [34]. Total cellular lipid content was strongly correlated with fenitrothion sensitivity, suggesting that lipophilic compounds will be more toxic to phytoplankton species with larger lipid fractions [35]. Fenitrothion at 5 and 10 mg/L caused bleaching of the chlorophyll pigment in several bluegreen algal species [19].

A study of the effect of dimethoate and thimeton on Sce-nedesmus incrassatulus revealed that growth was completely inhibited at concentrations above 0.075 and 0.5 mg/L, respectively. Both pesticides caused a considerable decrease in the level of algal proteins and carbohydrates. At concentrations greater than 1%, these pesticides significantly reduced cell density. Their toxicity resulted from a depletion of cell ca-rotenoid and, indirectly, of chlorophyll content [36]. Another study determined 10-d LC50s for Chlorella vulgaris and Anabaena doliolum exposed to dimethoate of 51 and 28.5 mg/L, respectively [24].

The organophosphate insecticide temephos stimulated growth of nitrogen-fixing bluegreen algae and the green alga Chlamydomonas at 10 and 100 μg/L [37]. The same concentrations, however, inhibited growth of the diatom Navicula pelliculosa and the green alga Chlorella pyrenoidosa [38]. The diatom Navicula minima and the bluegreen algae Coccochloris peniocystis and Oscillatoria sp. were less sensitive, and growth of Chlorella vulgaris and the bacterium Vibrio natrigens were not affected at temephos concentrations <5 mg/L [38].

The toxicity of seven organophosphate insecticides and their metabolites were determined using the Microtox® bio-assay (Azur Environmental, Carlsbad, CA, USA [39]). This assay utilizes the bioluminescent marine bacterium (Vibrio fisheri) as the test organism. An EC50 is determined using the metabolic indicator of reduced light output relative to the control. The EC50s of 5.2, 3.2, and 4.8 mg/L were determined for fonofos and its metabolites, methyl phenyl sulfone and thiophenol, respectively. Parathion had an EC50 of 8.5 mg/L, and its metabolite 4-nitrophenol had an EC50 of 13.7 mg/L. The EC50s of 46.3 and 18.6 mg/L were determined for chlor-pyrifos and its metabolite, 3,5,6-trichloro-2-pyridinol, respectively. The EC50s of 97.8, 213.9, and 5.6 mg/L were determined for isofenphos and its metabolites, salicyclic acid and isopropyl salicylate, respectively. Diazinon had an EC50 of 10.3 mg/L, and its metabolite hydroxypyrimidine had an EC50 of 886.4 mg/L.

Chlorpyrifos

Chlorpyrifos, like other organophosphorous insecticides, is an acetylcholinesterase inhibitor. The compound forms a reversible complex with the enzyme, disrupting proper nerve function [4]. Chlorpyrifos was toxic to the marine diatoms Skeletonema costatum and Minutocellus polyorphus at 0.24 and 0.64 mg/L (48-h growth rate EC50s), respectively [40]. Chlorpyrifos at 1 to 10 mg/L depressed growth in the bluegreen alga Anabaena sp. [19].

Other studies have examined chlorpyrifos toxicity in aquatic communities. Mani and Konar [41] found that 0.02 mg/L chlorpyrifos (dosed for a maximum of six times at an interval of 15 d) significantly reduced dissolved oxygen and increased free carbon dioxide. Zooplankton abundance was significantly reduced, whereas the phytoplankton remained unaffected. Butcher et al. [42] observed an increase in dissolved oxygen and a decrease in free carbon dioxide with chlorpyrifos concentration (4, 10, and 1,000 μg/L) in artificial ponds. Algal blooms were more prevalent in chlorpyrifos-treated ponds. Hurlbert et al. [43] reported an increase in phytoplankton abundance in freshwater ponds at 7.2 and 72 μg/L chlorpyrifos. Blooms of cyanobacteria (Anabaena, Anabaenopsis) occurred at both concentrations tested. Chlorophyll a content increased significantly 30 to 50 d after exposure in ponds dosed with 10 p,g/L chlorpyrifos [44]. Chronic exposure of Anabaena sp. to concentrations of 0.1 μg/L chlorpyrifos and 5 μg/L atrazine were conducted in indoor freshwater microcosms. No chlorpyrifos effects on phytoplankton were detected, whereas a small decrease in photosynthetic activity was indicated in the atrazine exposure [45]. The effect of chlorpyrifos in combination with nutrient loading was studied in indoor microcosms simulating drainage ditches. The combination of insecticide and nutrients increased phytoplankton abundance and periph-yton biomass to a greater extent than that observed with nutrient additions alone. The increase was thought to stem from fewer arthropod grazers [46,47]. In another study, 10 μg/L chlorpyrifos significantly reduced ciliate abundance of a natural microbial community. An increase in bacterial abundance and productivity and a decrease in chlorophyll a were also observed at 10 μg/L chlorpyrifos [29]. At a normal application concentration of 1.2 μg/L, chlorpyrifos caused persistent (up to 17 d following the initial application) reduction in growth of most phytoplankton in a freshwater pond community [48]. In general, these community study data indicate that release from zooplankton predation often leads to an increase in algae following treatment with organophosphorus pesticides.

Pyrethroid insecticides

A few studies have examined the effect of pyrethroid insecticides to phytoplankton. For example, the pyrethoid cy-fluthrin was tested at concentrations ranging from 2.5 to 62.5 g/hectare in natural and artificial ponds [49]. Fenvalerate (a synthetic pyrethroid insecticide) was examined by Day et al. [50]. No lasting effects on the phytoplankton were observed in any of the studies, and temporary changes in abundance were attributed to declines in zooplankton abundance.

Carbamate insecticides

Carbaryl had an EC50 of 5.0 mg/L, and its metabolite 1-naphthol had an EC50 of 3.7 mg/L when determined with the Microtox bioassay using the marine bacterium V. fisheri [39]. Carbofuran had an EC50 of 20.5 mg/L, while its metabolites carbofuran phenol and methylamine had EC50s of 60.9 and 34.6 mg/L, respectively [39]. Peterson et al. [51] examined the interspecific sensitivity of Scenedesmus quadricauda and Selenastrum capricornutum (green algae), Nitzschia sp. and Cyclotella meneghiana (diatoms), and Microcystis aeruginosa, Oscillatoria sp., Pseudoanabaena sp., Anabaena in-aequalis, and Aphanizomenon flos-aquaeb (cyanobacteria) to 3.7 mg/L carbaryl (an expected environmental concentration [EEC]) and found >50% inhibition of 14C uptake in 90% of the algae tested. In contrast, another carbamate insecticide, carbofuran, had relatively low toxicity to most species tested when applied at the EEC of 0.67 mg/L [51]. Christie [30] found that 100 mg/L of the insecticide carbaryl reduced the population density of Chlorella pyrenoidosa by 30% and was inhibitory at concentrations as low as 0.1 mg/L. Marine phytoplankton were also susceptible to carbaryl, which was lethal to two species at 1 mg/L and to all five species tested at 10 mg/L [52]. In the bluegreen algae, carbaryl at 10 mg/L did not affect growth of Nostoc muscorum but enhanced the growth of Cylindrospermum [19].

Organotin insecticides

In an aquatic microcosm study, the organotin pesticide azo-cyclotin was found to inhibit picoplankton <2 μm and algae of 2 to 10 μm at ≥135 μg/L, and blooms of less sensitive species were observed (e.g., Kirchneriella sp.) [53]. The toxicity of tributyltin oxide and tributyltin chloride was examined with the marine diatoms Skeletonema costatum and Minutocellus polyorphus. The 48-h growth rate EC50s determined for tributyltin oxide were 0.34 and 0.33 mg/L for S. costatum and M. polyorphus, respectively. Similar values of 0.33 mg/L (S. costatum) and 0.36 mg/L (M. polyorphus) were determined for tributyltin chloride [40].

Herbicides

Only at high concentrations did the phenoxyalkane herbicide 2,4-D have adverse effects on algal populations [18]. A study by Faust et al. [31] found that reproduction of Chlorella fusca was not inhibited below 88.86 g/L of 2,4-D. Another study using four filamentous algal species showed no effect on chlorophyll production at ≤100 mg/L 2,4-D after 7 d [33]. A comparison of nine algal species exposed to the EEC of 2.9 mg/L 2,4-D found <10% inhibition of 14C uptake in all the algae tested [51]. Low toxicity to the same algal species was also observed with another pheoxyalkane herbicide, 2-methyl-4-chlorophenoxyacetic acid, tested at an EEC of 1,400 mg/L [51].

The toxicity of 2,4-D and its metabolites was determined with the Microtox bioassay using the marine bacterium V. fisheri [39]. The 2,4-D had an EC50 of 100.7 mg/L, whereas its hydrolysis metabolite 2,4-dichlorophenol had an EC50 of 5.0 mg/L. The structurally similar herbicide 2,4,5-T had an EC50 of 51.7 mg/L, and its metabolite 2,4–5-trichlorophenol had an EC50 of 1.8 mg/L.

In an ecological risk assessment of the brominated herbicide diquat, Campbell et al. [54] cited 7-d growth rate EC50 values for nine algal species, ranging from 1 μg/L for the pyrrophyte Peridinium cinctum to 2,940 μg/L for Euglena gracilis. Bluegreen algae and chrysophytes were more sensitive to diquat than the green algae [54]. A comparison of nine algal species exposed to EEC of 0.73 mg/L diquat found 53 to 69% inhibition of 14C uptake in the two green algal species, 99 to 100% inhibition in the two diatom species, and 100% inhibition in all five species of cyanobacteria tested [51]. Diquat significantly altered the algal and bacterial densities of naturally derived microbial communities tested in freshwater microcosms at concentrations greater than or equal to 0.3 mg/L [55]. There was also a significant reduction in species richness of protozoa at diquat concentrations <0.3 mg/L, with no observable recovery 21 d after dosing [55].

In contrast, another brominated herbicide, bromoxynil, did not significantly inhibit 14C uptake in any of the algal species when tested at the EEC of 0.28 mg/L [51]. In another study, paraquat application to naturally derived freshwater algal communities showed significant inhibition of the cyanobacteria (especially Phormidium) at 10 μg/L [56].

Peterson et al. [51] also tested two pyridine herbicides at their EEC: picloram (1.8 mg/L) and triclorpyr (2.6 mg/L). There was no significant inhibition of carbon uptake by these compounds in any of the algal species tested. The aldehyde herbicide acrolein, however, inhibited carbon uptake in all nine algal species by <95% at an EEC of 1 mg/L. Glyphosate (EEC of 2.8 mg/L), a glycine derivative herbicide, significantly affected carbon uptake in the diatoms Nitzschia sp. and Cyclo-tella meneghiana and the nitrogen-fixing cyanobacterium Aphanizomenon flos-aquae but had either no significant effect or a stimulatory effect in the other seven algal species. In another study, 4-h EC50 values ranging from 8.9 to 89 mg/L glyphosate were determined for freshwater periphyton communities [57].

Phenylurea herbicides have many significant effects on algae. Additions of monuron at 4 mg/L to cultures of Chlorella sp. caused growth rates to decline [58]. In a study by Ukeles [52], diuron was lethal to marine algal species at 4 μg/L and ranked as the most toxic phenylurea tested. In order of decreasing toxicity, the compounds were diuron < monuron < neburon < fenuron. The compounds reflect differences in photosynthesis inhibitory abilities and differences in water solubility, with low solubility causing accumulation in the algal cell. Another phenylurea compound, chlortoluron, was inhibitory of reproduction in Chlorella fusca at 23.39 mg/L [31]. Peterson et al. tested the toxicity of four sulfonylurea herbicides-hlorsulfuron(0.02 mg/L), thametsulfuron-methyl (0.015 mg/L), metsulfuron-methyl (0.003 mg/L), and triasul-furon (0.018 mg/L)-singn ne algal species [51]. At the EEC applied, there was little or no inhibition of carbon uptake in any of the algal species tested. In contrast, the substituted urea herbicide tebuthiuron was very toxic to the same algal species. Significant inhibition of 14C uptake was observed in all species tested at an EEC of 5.9 mg/L tebuthiuron [51]. An acetanilide herbicide and an imidazolinone herbicide (metolachlor and imazethapyr, respectively) were also tested but exhibited little toxicity to the algal species. Of the 9 algal species tested, metolachlor (EEC of 3 mg/L) significantly inhibited carbon uptake in the green alga Selenastrum capricornutum and the cyanobacterium Pseudoanabaena sp., while imazethapyr (EEC of 0.067 mg/L) only caused significant inhibition in the cyanobacterium Microcystis aeruginosa [51].

Chloroplast mutations may lead to the loss of herbicide binding and inhibition in selected tolerant species. Diuron resistant strains of Chlamydomonas reinhardii, Bumilleriopsis filiformis, and Euglena gracilis have been discovered. The diruon-resistant C. reinhardii line was less resistant to the photosynthesis inhibitors simazine and atrazine than to diuron and monuron but normally sensitive to bromacil. The diuron-resistant E. gracilis line is also resistant to o-phenanthrolene [31].

The triazine herbicide anilazine inhibited reproduction of Chlorella fusca at 1,389 mg/L, whereas simazine had the same effect at only 26.14 mg/L [31]. Simazine (1 mg/L) significantly reduced chlorophyll production in three filamentous algal species after 7 d [33]. Another study compared the toxicities of two triazine compounds on freshwater algae. The triazine gar-doprim decreased growth rates of a freshwater algal community above 0.05 mg/L, whereas gesapax exhibited no harmful effects at 0.1 mg/L [59]. Peterson et al. [51] examined the interspecific sensitivity of Scenedesmus quadricauda and Selenastrum capricornutum (green algae), Nitzschia sp. and Cy-clotella meneghiana (diatoms), and Microcystis aeruginosa, Oscillatoria sp., Pseudoanabaena sp., Anabaena inaequalis, and Aphanizomenon flos-aquae (cyanobacteria) to five triazine herbicides applied at the EECs. Atrazine (2.67 mg/L), cyan-azine (2.67 mg/L), hexazinone (2.87 mg/L), metribuzin (2.67 mg/L), and simazine (2.67 mg/L) inhibited carbon uptake of all algae by more than 50%.

A single-pulse, 21-d exposure of a naturally derived algal community to the chloroacetamide herbicide alachlor reduced chlorophyll a levels and altered taxonomic composition at ≥ 10 μg/L, whereas concentrations ≥30 μg/L significantly reduced total algal cell densities [60]. Alachlor and atrazine were found to act additively on benthic algal communities in artificial streams. Algal community biovolume was reduced at 90 μg/L alachlor and at 12 and 150 μg/L atrazine [61]. A 96-h growth rate EC50 of 5.7 mg/L alachlor was determined for the es-tuarine phytoplankton Pavlova sp. [62]. A 96-h mixture EC50 of 64 μg/L for atrazine and of 2,126 μg/L for alachlor was also determined for Pavlova sp., which was a slightly greater than additive effect [62].

Atrazine

Atrazine is an s-triazine herbicide used primarily to control broadleaf plants and grassy weeds. Atrazine is algistatic, inhibiting photosynthesis by blocking electron transport during the Hill reaction of photosystem II [63,64]. Algal responses to atrazine vary widely, depending on concentrations used, duration of exposure, and algal species tested. Many studies have examined the effects of atrazine on phytoplankton (see reviews by Huber [65] and Solomon et al. [66]). Some algal toxicity results from atrazine studies are summarized in the following.

Stratton [67] found growth rates to be suppressed from 100 to 5,000 μg/L atrazine for five different algae. Mayasich et al. [68] reported Phaeodactylum tricornutum to be unaffected by 50 μg/L atrazine, while the same concentration of atrazine inhibited the growth rate of Nannochloris oculata by about 35%.

Decreased chlorophyll content (41–67%) was observed at 1 μg/L atrazine in the green algae Chlorella vulgaris and Stigeoclonium tenue and in the bluegreen alga Oscillatoria lutea after 7 d of exposure [33]. Abou-Waly et al. [69] determined 3-, 5-, and 7-d growth rate EC50s of 58, 469, and 776 μg/L, respectively, for the bluegreen alga Anabaena flos-aquae. Parrish [70] reported a decrease in culture growth and fluorescence at 13 to 22 μg/L atrazine for the saltwater phy-toplankter Skeletonema costatum after 5 d of exposure. Walsh et al. [40] reported 48-h EC50 values of 50 and 20 μg/L for the marine diatoms Skeletonema costatum and Minutocellus polyorphus, respectively. Toxicity tests with five species of the green algae Scenedesmus sp. yielded 3- to 14-h growth rate EC50s of 21 to 300 μg/L [67,71]. Atrazine toxicity was compared using eight individual algal species in microcosm and experimental pond tests [72]. Mean species 24-h EC50 values for 14C uptake ranged from 37 to 308 μg/L, with green algal species being more sensitive than bluegreens. Microcosm EC50 values ranged from 103 to 159 μg/L, and the EC50 determined from the experimental ponds was 100 μg/L [72].

Another study compared the toxicity of atrazine, isopro-turon, and mecoprop to the green alga Scenedesmus subspi-catus. The 96-h growth rate EC50s determined were 21 μg/L atrazine, 21 μg/L isoproturon, and 102,660 μg/L mecoprop [73].

A 28-d atrazine exposure of four green algae (Chlamydo-monas sp., Chlorella sp., Pediastrum sp., and Scenedesmus quadricauda) and four diatoms (Cyclotella gamma, Cyclotella meneghiniana, Synedra acus, and Synedra radians) yielded EC50s ranging from 27.6 to 110.6 μg/L for the green algae and 88.9 to 429.7 μg/L for the diatoms [74]. In another study, growth was inhibited approx. 85% in the green alga Chla-mydomonas reinhardtii by 0.23 μM atrazine, whereas growth was inhibited 60% in Scenedesmus quadricauda at the same concentration [75]. A laboratory study with the estuarine phy-toplankton Pavlova sp. determined a 96-h growth rate EC50 of 147 μg/L atrazine [62].

Mayasich et al. [68] reported Phaeodactylum tricornutum to be unaffected by 50 μg/L atrazine, while the same concentration inhibited the growth rate of Nannochloris oculata by about 35%. The green alga Chlamydomonas sp. was found to be quite sensitive, with a 1-h EC50 of 60 μg/L using oxygen production inhibition as an end point [76]. One-hour EC50 values for eight species of marine diatoms ranged from 84 μg/L for Cyclotella nana to 460 μg/L for Navicula inserta [76]. In a 7-d chronic atrazine exposure, Plumley and Davis [77] reported reduced rates of photosynthesis and reduced cell numbers for diatom species isolated from a Georgia salt marsh (Thalassiosira fluviatilis and Nitzschia sigma). Based on the least-effect level of atrazine to diatoms, the maximum safe level determined for atrazine in the salt marsh was 10 μg/L.

In a study of atrazine effects on natural plankton assemblages, 20 μg/L reduced algal productivity and biomass, followed by recovery to control levels after 7 d [78]. Hamilton et al. [79] observed significant reductions in phytoplankton species diversity at 100 μg/L atrazine applied to lake enclosures. The changes in community composition persisted for 77 d. Net primary productivity, pH, and net productivity/respiration ratios were reduced in naturally derived algal communities at 100 to 200 μg/L atrazine [80]. In another study, atrazine (153 μg/L) reduced chlorophyll levels, followed by recovery between 7 and 14 d. Higher levels remained inhibitory to primary productivity and chlorophyll levels after 14 d and resulted in a decline in green algal colonies [81]. A microcosm study of periphyton communities found that atrazine concentrations of 10 μg/L or less had a stimulatory effect on protein biomass, chlorophyll, and species richness, whereas higher concentrations led to a significant loss of benthic algal species and biomass and lower net oxygen production [82]. It was also found that atrazine concentrations of 0.8 to 1.56 μg/L reduced productivity of attached lake algae by 21 to 82% [83]. An atrazine study in flow-through wetland mesocosms revealed that periphyton net primary productivity was significantly depressed at ≥25 μg/L (9–27-d exposure). Periphyton developed resistance to atrazine at concentrations of 50 μg/L or higher [84]. In pond microcosms receiving continuous atrazine doses at concentrations of 0.5 to 5,000 μg/L, Brockway et al. [85] measured a reduction in primary production at ≥50 μg/L.

Studies have shown that periphyton communities do not necessarily become more resistant to atrazine after previous exposure [56,85–88]. In some cases, algae actually exhibited increased sensitivity to the herbicide in an acute exposure following a chronic exposure and recovery period [62,86]. Nys-trom et al. [87] also found low potential for algae to become tolerant to atrazine using a freshwater periphyton community.

Changes in community composition were reported in many studies, but the relative sensitivity of different taxa was not consistent. In pond mesocosm studies, planktonic green algae and flagellates were reduced while cryptophytes (especially Cryptomonas) and chrysophytes (especially Mallomonas) increased in abundance [78]. Hamilton et al. [79] measured a reduction in green algae, diatoms, and dinoflagellates in enclosed phytoplankton communities exposed to 100 μg/L atrazine, whereas chrysophytes were unaffected and cryptophytes increased slightly. Cryptophytes and diatoms were reduced in laboratory streams exposed to 100 μg/L atrazine, whereas bluegreen algae increased [86]. A similar taxonomic shift was observed in estuarine mesocosms dosed with 40 and 160 μg/L atrazine [89]. In another study, 100 μg/L eliminated blue-green algae but had little effect on diatoms [90].

Reductions in dissolved oxygen concentrations are commonly observed with atrazine-induced reductions in primary productivity [29,85,91]. Other water quality changes include reductions in pH and inorganic carbon and increases in alkalinity and conductivity. These responses are related to reduced photosynthetic uptake of bicarbonate. Several investigators have reported increases in inorganic nitrogen [81,84,85,92] or phosphorus [81,84,93] due to reductions in nutrient uptake rates.

The most toxic metabolite of atrazine is deethylatrazine [67]. Deethylatrazine was found to significantly reduce chlorophyll a content, phototrophic carbon assimilation, photo-trophic biovolume, and dissolved oxygen levels of estuarine microbial communities at 50 μg/L [29]. No significant effects on bacterial abundance, productivity, or heterotrophic protozoan abundance were observed in that study.

Fungicides

In a comparison of thiocarbamates, maneb was more inhibitory than zineb to the growth of Euglena gracilis, while nabam was toxic at all levels tested [94]. Bluegreen algae are fairly resistant to insecticides but are particularly sensitive to fungicides mentioned previously [19]. Ethyl mercury phosphate was lethal to all marine phytoplankton species tested when incorporated at a level of 60 μg/L in the culture media [52]. Harris et al. [95] found that three organomercury fungicides, at less than 1 μg/L, reduced growth and photosynthesis in phytoplankton. Data such as these revealed that marine and freshwater phytoplankton were sensitive to organomercury compounds at levels below those proposed for water quality standards at the time, suggesting that entry of such compounds into natural waters should be prevented. A more recent study [51] compared the sensitivity of nine algal species to the tri-azole derivative fungicide propiconazole. At the EEC of 0.08 mg/L, inhibition of 14C uptake ranged from 0 to 31%, with the green algae Scenedesmus quadricauda and the cyanobac-terium Microcystis being the most sensitive species tested.

ECOLOGICAL RELEVANCE OF PESTICIDE TOXICITY TO MICROORGANISMS

It is evident that there are considerable differences in pesticide sensitivity among microbial species and that the use of an uncertainty factor is necessary to provide an acceptable margin of safety in evaluating the hazard presented by these chemicals to aquatic environments.

Using the microbial toxicity data summarized here and environmental monitoring data of pesticide concentrations, we can predict which pesticides are most likely to cause an effect in aquatic ecosystems. Some of the pesticides described here are no longer used in the United States but are still routinely detected in surface waters (e.g., DDT). These chemicals may pose a threat to microorganisms when they become resus-pended in marine and estuarine sediments because of tidal action or dredging.

The lipophilic nature of many pesticides may pose a threat to higher estuarine organisms. For example, although endo-sulfan was not toxic to algae at levels likely to be found in the environment (< 1 μg/L), the compound may bioaccumulate in algae and be consumed in higher concentrations by grazers. Rao and Lal [26] found rapid uptake of the commonly used agricultural pesticide endosulfan in the cyanobacteria Aulosira and Anabaena, with levels reaching 700 times the exposed dose within 48 h. The metabolite endosulfan sulfate bioac-cumulates more and is more persistent (half-life on the order of months) than the parent isomers [96]. The amount of pesticide accumulated depends not only on adsorptive and transfer processes but also on the amount continued to be released into the water and transformation of the chemical [97]. There is, therefore, the potential for chronic effects, such as evolved sensitivity, bioconcentration, and impacts on higher trophic levels. This is especially true in estuaries, where contaminated sediments are continually resuspended because of tidal action and dredging. Pesticides accumulated in prey species may then cause toxicity in the organisms consuming them or continue to bioconcentrate through the food web. Use of environmentally persistent pesticides in the southeastern U.S. coastal region should continue to be carefully monitored.

Atrazine is one of the most extensively used herbicides in the United States and is routinely detected in aquatic habitats because of runoff from agricultural fields [98,99]. Atrazine accounts for 60% of the total pesticide volume applied to crops each year, with an estimated 29 million kg of atrazine active ingredient applied in 1989 alone [100].

An ecological risk assessment of atrazine was conducted by Solomon et al. [66]. They concluded that although atrazine effectively impairs phytoplankton populations at concentrations typically found in the environment, the ecological system usually recovers quickly from such exposures. Ecologically significant effects were considered to occur only at concentrations of 50 μg/L or greater. It was further recommended that site-specific risk assessments be conducted for aquatic systems where atrazine is heavily used. The EC50s summarized for marine phytoplankton ranged from 20 to 600 μg/L. In a literature review of atrazine toxicity in aquatic ecosystems, Huber [65] concluded 20 μg/L to be the no-observed-effect concentration. Studies have shown, however, that growth of phytoplankton can be inhibited at concentrations as low as 0.1 μg/L [101,102].

Surface water quality monitoring of 149 sites in 122 mid-western river basins indicated that 52% of the sites exceeded the U.S. EPA's maximum contaminant level of 3.0 μg/L for atrazine and that 32% of the sites exceeded the maximum contaminant level for alachlor of 2.0 μg/L [103]. Maximum atrazine concentrations were >100 μg/L, and maximum alachlor concentrations were >55 μg/L. An estuarine monitoring study of atrazine concentrations along the mid-Texas coast, USA, found atrazine contamination to be quite pervasive [62]. Greater than 97% of samples collected had a detectable level of atrazine (February-October 1993). Additionally, 52% of samples collected had atrazine concentrations exceeding 1 μg/L, and over 5% of the samples collected along the mid-Texas coast had atrazine concentrations exceeding the 20-μg/L no-observed-effect-concentration value identified by Solomon et al. [66] and Huber [65]. It was concluded that phytoplankton, which are chronically exposed to atrazine along the mid-Texas coast may not be able to recover because of the ubiquitous nature of the contaminant [62]. In addition to atrazine, eight other herbicides (cyanazine, hexazinone, metribuzin, simazine, acrolein, diquat, carbaryl, and tebuthiuron) were found to be highly phytotoxic to algae when tested at the EECs [51]. Based on the data available, herbicides have high potential to elicit significant effects on aquatic microorganisms at environmentally relevant concentrations.

The toxicity of pesticides to microorganisms may result in a reduced food source for higher trophic levels. For example, abundance of a dominant zooplankton species was reduced by 75% in ponds treated with 500 μg/L atrazine [78]. Compositional changes in the phytoplankton and decreased photo-synthetic biomass and productivity have been shown to cause long-term reductions in the food resources of certain grazers [1]. These changes also affect the quantity and quality of nutrients cycled through the microbial food web and passed to higher trophic levels [104].

Also, microbes may be affected indirectly by pesticide effects occurring at higher trophic levels. For instance, phytoplankton populations increased after fenvalerate (a synthetic pyrethroid insecticide) caused a significant decline in the population of large-bodied cladocerans [50]. Similarly, sublethal concentrations of endosulfan inhibited filtration and ingestion rates of Daphnia magna fed the unicellular alga Nannochloris oculata. Filtration and ingestion rates were reduced to 50% of the control values at 0.44 and 0.61 mg/L endosulfan, respectively [105]. Significant decreases in rates of filtration and ingestion were also observed with the rotifer Brachionus ca-lyciflorus [106].

AREAS NEEDING MORE RESEARCH

Many questions remain regarding what levels of pesticides will be toxic to marine and estuarine microorganisms. In addition, pesticide data are clearly lacking for protozoa. The toxicity of pesticides to this important aquatic group has been generally overlooked. Bacterial studies are relatively abundant; however, most deal with degradation of pesticides by soil bacteria rather than toxicity of pesticides to aquatic bacteria. Alteration of bacterial growth and/or productivity by pesticides has the potential to affect other microbial components as well. Future research should assess the toxicity of pesticides to the microbial food web as a whole.

Another point for consideration is the influence of water quality on the toxicity of pesticides to microorganisms. Factors such as pH, microbial cell density, salinity, nutrient concentrations, and chemical form of the pesticide in the system may alter microbial response to the pesticide. For instance, agricultural pesticides are usually applied in conjunction with fertilizers. Nutrient additions can cause stimulation or inhibition of photosynthesis and respiration, either of which can alter pesticide toxicity. Studies comparing pesticide responses under nutrient-enriched versus nutrient-limited conditions would be useful contributions to the literature. For example, there is evidence that atrazine becomes more toxic to phytoplankton under nutrient-enriched conditions [78; DeLorenzo et al., in preparation]. Another study found that natural periphyton communities in low-nutrient treatments were less likely to recover from herbicide stress than those in medium- and high-nutrient treatments [107].

Testing conditions also play a role in the toxicity observed and should attempt to simulate the ecosystem as much as possible. For example, three weeks after treatment with the herbicide diquat (0.04–21.0 mg/L), microbial communities in microcosms without sediment still exhibited decreased structure and function, whereas in microcosms amended with natural lake sediments, dosed communities were not significantly different from controls by two weeks postdose [108].

It is also important to remember that aquatic systems may be contaminated with a number of different pesticides. This is especially relevant when predicting how pesticide effects observed in the laboratory compare to responses occurring in the environment. For example, interactions between combinations of organochlorine compounds and polychlorinated bi-phenyls have been documented. The DDT metabolite DDE and polychlorinated biphenyls were far more inhibitory to a marine diatom, Thalassiosira pseudonana, in combination than they were individually [109]. More studies are needed involving pesticide mixtures and microorganisms.

Lastly, the toxicity of pesticide degradation products is a major area requiring further study. Many of these compounds linger in aquatic systems long after the parent compound is gone. For example, deethylatrazine is the most commonly detected transformation product of atrazine in aquatic ecosystems [110,111]. Deethylatrazine has lower Kd and Koc values than atrazine and is therefore more readily transported in the aqueous phase. It is also persistent in surface waters. Pereira and Hostettler [99] found no evidence of degradation of deethy-latrazine during its transit time (45–65 d) in the entire navigable reach of the Mississippi River, USA. A laboratory study found this degradation product to be nearly as toxic as the parent compound to estuarine microbial communities [29].

As our coastlines continue to become more developed and populated, the presence of pesticides in marine and estuarine environments is likely to increase. To successfully protect and manage our coastal resources, more information is needed regarding the hazard of pesticides to aquatic microorganisms forming the base of the food web.

Acknowledgements

The National Ocean Service (NOS) does not approve, recommend, or endorse any proprietary product or material mentioned in this publication.

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