Open Literature Review Summary

Chemical Name: Atrazine

CAS No: 1912-24-9

ECOTOX Record Number and Citation:

Johnson, B.T. 1986. Potential impact ofselected agricultural chemical contaminants on

a northern prairie wetland: A microcosm evaluation. Environ. Toxico!. Chem. 5:473-

485. (mt.io qt5''l',~ ~D?»)

Purpose of Review:

Determine potential effects of atrazine on an aquatic community. Endpoints from this

study were also used to parameterize the Comprehensive Aquatic System Model (CASM)

and establish the model-associated LOC for atrazine.

Date of Review:

January 14,2009

Purpose of Study:

This study examines the potential effects of atrazine to aquatic plants and animals in

laboratory microcosms.

Summary of Study Findings:

Methods: This study examined the effects of 3 different herbicides (including atrazine)

and 3 different insecticides, in a nested experimental design. Also, some animal toxicity

experiments were conducted to examine the effects ofthe insecticides in particular. In

this review, only the methods pertaining to atrazine effects on the microcosms will be

fully described; methods not relevant to atrazine will be omitted, and methods

tangentially relevant to atrazine or conducted without the microcosms will be briefly

summarized.

Experiments were conducted in microcosms consisting ofsediment, water, plants (algae

and macrophytes), and animals (planktonic and benthic invertebrates) in 4 L glass jars.

Each jar was filled with 3.8 L of water and hydrosoil (sediment) at a ratio of9:1 by

volume, and covered with a circular glass plate (15 cm diameter) to reduce water loss.

Each plate had a central hole, about 5 mm in diameter, for gas exchange. Microcosms

were randomly located on a bench in a 7 ft x 15 ft x 7 ft environmental chamber (Panel

'••111.1.1

2087856
Key, Earth City, MO) maintained at 20±1°C, with a ceiling bank of t1uorescent lights

(Gro-Iux) providing 1,400 lux in a 16 hour light8 hour dark photoperiod. These

conditions were called "standard conditions".

Soil matter (sediment from a permanent wetland and topsoil from an adjacent agricultural

field) came from Clay County Waterfowl Production Area, about 21 km southeast of

Vermillion, South Dakota. Sediments, portions of overlying water, and field soil were

removed in the spring and fall and transported to the laboratory (Columbia National

Fisheries Research Laboratory [CNFRLJ) within 48 hours. Topsoil contained 2.6%

organic matter and 0.22% Kjeldahl nitrogen, and wetland sediment contained 9.8%

organic matter and 0.79% Kjeldahl nitrogen. Soybeans and corn were grown in the fields

around the wetland site, but gas-liquid chromatographic-mass spectrophotometric

analyses ofthe wetland samples found no detectable pesticide residues.

Microcosm water was a mixture ofstandard reconstituted water (EPA, 1975, Alethodfor

acute toxicity tests with fish, macroinvertebrates) and amphibians) and natural wetland

water in a ratio of 8: I by volume. Reconstituted water had the following amounts ofsalts

per 10 L: 0.48 g NaHCOJ, 0.30 g CaS04, 0.30 g MgS04, and 0.02 g KCI. The pH,

alkalinity, and conductivity were 7.2 and 35 mg/L (as CaC03) and 150 Ilmhos/cm,

respectively. Water loss due to sampling was replenished with reconstituted water, but

replenishments never exceeded 0.5% (v/v) ofthe total unit volume. Evaporative water

loss was replaced weekly with deionized water from a Millipore reverse osmosis unit

(Millipore Corporation, Bedford, Massachusetts), which had pH 6.0, alkalinity and total

hardness 10 and 1.0 mg/L (as CaCOJ), and conductivity about 1.0 Ilmhos/cm.

After allowing one week for the sediments to settle and the water to clear, three types of

macrophytes were added to each microcosm: Lemma sp. (duckweed, surface floating, 10

plants of ~50 mg each), Ceratophyllum sp. (coontail, nonrooting, submergent, 15 cm

segment of 109), and Elodea sp. (American Elodea, rooting, submergent, 15 cm segment

of 5 g). Elodea plants were weighted with a small segment of glass tubing to hasten

rooting in the sediment. Natural communities of invertebrates and algae then developed

within each microcosm, with patches of bluegreen and filamentous green algae visible on

the water surface within 1 week. Cladocerans, ostracods, copepods, rotifers, and tubedwelling chironomids (attached on jar sides) appeared within several weeks. Before

treatments, microcosms were cross-seeded to attain uniform ecosystem compositions.

Microcosms were allowed to equilibrate for about 2 weeks before treatment with the

pesticides. (Timeline is summarized as in the article; it is unclear from when the 2 week

equilibration time is counted.)

Pesticides applied in the experiment were: atrazine, trifluraIin, triallate, carbofuran,

[onofos, and phorate (the first three being herbicides and the last three being

insecticides). Atrazine was Aatrex, 85.5% active ingredient wettable powder from Ciba~

Geigy Corporation, with a solubility of70 mg/L at 20°e. Each ofthe six pesticides were

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tested at three different concentrations, 10, 100, and 1,000 flg/L, which were introduced

by stock solutions in acetone or water based on % active ingredient, mixing with 1.0 g of

topsoil in alSO m1. beaker and allowing to sit for one hour, adding 50 mL reconstituted

water, and then gently stirring with a glass rod and slowly pouring the total slurry into the

water surface ofthe microcosm. The microcosm water was not mixed. Control

treatments received the soil and carrier solvent. Treatments were made once and not

repeated, and the experiments were terminated after 30 days. (No atrazine concentration

measurements were taken in the microcosms.)

Each microcosm series consisted offour treatments with four replicates per treatment,

with one treatment being the control. All measurements ofspecific variables were taken

in triplicate unless otherwise stated. Most samples were taken weekly. Data were

analyzed in a split-plot design, in which the linear statistical model contained the main

effect at each treatment, replicates within treatment, weeks, and the interaction of

treatment x week, using ANOVA as implemented in SAS. Within-treatment replicates

were used as errors to test the treatment effect, and the residual mean square was used as

the error to test the effects of week and treatment x week. Simple one-way ANOVA was

used to analyze variables measured only at the end ofthe experiment. Ifthe overall F

value was significant at p :s 0.05, Fisher's least-significant-difference test was used to

find mean differences. Samples were tested for homogeneity of variance.

48-hour EC50 values were determined for the insecticides using standard acute toxicity

tests with Daphnia magna and Chironomus riparius; for the herbicides, published EC50

data were confirmed by performing bracketing experiments above and below the reported

EC50. Additionally, tests were performed to evaluate direct pesticide effects on daphnid

growth and reproduction: a microcosm acute toxicity test (14 or 30 days) with first-instar

«24 hours old) D. magna and early fourth-instar C. riparius larvae using standard

toxicity test methods; a viability test wherein 25 mature (>2,500 flm long), gravid

daphnids were added to each microcosm 48 hours before treatment, with the population

monitored throughout the study and replenished when less than 5 organisms were

observed; and a 7-day chronic toxicity test (not in microcosms). (Further details on these

experiments will not be summarized here, but are in the article.)

Phytoplankton toxicity tests assessed the short-term growth inhibition of Selenastrium

capricornutum in microcosm water. Actively growing algae were concentrated by

centrifugation and resuspended, and 1 mg (dry weight) ofthese cells was placed into a

125 mL Erlenmeyer flask containing 50 mL of a microcosm's water. An algal bioassay

was conducted according to the methods of a 1978 EPA report on the protocol for bottle

tests with this alga (EPA-600/9-78-0 18). The flasks were harvested after 7 days and the

dry weight biomass ofthe algal cells was determined gravimetrically with tared Gelman

glass fiber filters.

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Water quality measurements (dissolved oxygen, conductivity, pH, hardness, and

alkalinity) and total organic content ofthe sediments were determined using American

Public Health Association (1980) standard methods. Total nitrogen and total phosphorus

concentrations in the water and sediment were measured with a Technicon Auto Analyzer

II (Technicon Inc., Tarrytown, New York) by a colorimetric method after persulfate

digestion.

Gross primary productivity and community respiration wcre estimated from diurnal

changes in dissolved oxygen in the microcosms, over two days, following the methods of

McConnell (1962, Productivity relations in carboy microcosms, Limnol. Oceanogr.).

The glass cover of the microcosms was removed 1 day before and during sampling days.

The three-point diurnal measurements were made weekly.

Total macrophyte biomass was measured at 30 days. All macrophytes were collected

from each microcosm, washed to remove soil, dried for several days at 50°C, cooled, and

weighed, to determine total macrophyte dry weight biomass.

Sediment microbial activity bioassays were conducted by removing 25.0 g (wet weight)

of hydrosoil and 25 mL of water from a microcosm and mixing in 1,000 Ilg/L pesticide

with a magnetic stirrer, with tests beginning after I hour of incubation at 20°e. Further

tests at 100 and 10 Ilg/L were only to be conducted ifthe initial tests showed at least a

25% deviation from controls. Respiration (oxygen consumption), electronic transfer

system (ETS) activity, monophosphotase activity, and mineralization of glucose were

tested using published methods. Oxygen readings were taken every 15 minutes for I

hour with a Gibson differential respirometer using a constant pressure manometric

technique. ETS activity was measured with the 2-(p-iodophenyl)-3-(p-phenyl-5-phenyl)

tetrazolium chloride (INT) method, with an INT mixture (6.0 mL microcosm water, 2.0

mL 0.2% INT, 1.0 mL 0.1 % sodium citrate) blended into I mL ofsediment slurry in a 50

mL centrifuge tube with a vortex mixer, the mixture incubated under nitrogen gas in a

stainless steel anaerobic container for 1 hour at 28°C, and the INT-tormazan content of

the solids measured as follows. The INT-formazan was extracted with 25 mL 95%

ethanol in a vortex mixer until no red-colored formazan was being removed, the ethanol

batches were mixed, and an aliquot was centrifuged at 10,000 g at 3°C for 5 minutes and

analyzed with a Spectronic 100 spectrophotometer with a flow-through cell (Bausch and

Lomb, Rochester, New York) at 480 nm. Absorbance values were corrected with

controls and compared to an INT-formazan standard curve. Monophosphotase activity

was measured with a modifiedp-nitrophenol (PNP) method, with a PNP mixture (7.0 mL

buffer of 0.05 M tris(hydroxymethyl)-aminomethane [THAM] at pH 8.0 and 2 mL of 1%

PNP) added to 1 mL ofsediment slurry in a 50 mL centrifuge tube, the mixture incubated

at 35°C for 1.5 hours while mixing every 15 minutes on a vortex stirrer, and the PNP

content of the solids measured analyzed as follows. The uptake was terminated by

adding 10 mL of a chilled acetone:THAM solution (l: 1 by volume, pH 12) and placing

the samples on crushed icc, tben the samples were centrifuged at 10,000 g and 3°C tor

about 5 minutes and spectrophotometrical1y analyzed at 410 nm. Absorbance values

4
were corrected to controls (types of controls were hydrosoil with substrate, which were

not incubated, and hydrosoil without substrate, which were incubated). Glucose uptake

was measured using a 14C-labeled glucose (uniformly labeled, 300 mCi/mM, Pathfinder,

Inc., 81. Louis, Missouri). ] mL of sediment slurry was added to a sterile 50 mL

Erlenmeyer flask containing 10 mL of microcosm water, 0.1 ).lCi of 14C-Iabeled glucose,

and 25 ).lg nonlabeled glucose, and the flasks were tightly sealed with a rubber stopper

and incubated for] hour at 20°C in the dark. Labeled CO2 (corresponding to mineralized

glucose) was captured in a 1.0 mL volume of 0.2 N KOt! in a suspended glass vial, the

reaction was stopped by acidifying the mixture with 0.1 N HCl and placing on crushed

ice, and samples were refrigerated overnight at 2°C for increased recovery. The total

volume ofthe KOH trapping solution was analyzed with liquid scintillation counting,

after mixing with a 15 mL Fluoralloy:Triton X-I 00 scintillation cocktail (14: 1 by

volume) (cocktail and scintillation counter by Beckman Instrument Inc., Fullerton,

California). Values were corrected for counting efficiency, quench, and background

radiation.

Results: This study examined the effects of 3 different herbicides (including atrazine)

and 3 different insecticides, in a nested experimental design. Also, some animal toxicity

experiments were conducted to examine the effects ofthe insecticides in particular. In

this review, only the results pertaining to atrazine effects on the microcosms will be fully

described; results not relevant to atrazine will be omitted, and results tangentially relevant

to atrazinc or conducted outside the microcosms will be briefly summarized.

The published EC50 values for atrazine toxicity to D. magna and C. riparius were

confirmed; EC50 values were 3,600 and 1,000 ).lg/L, respectively. Other animal

experiments found no evidence of atrazine influence on daphnid survival, growth, or

reproduction (same population of] was established posttreatment in all treatment levels,

including controls), no acute toxicity effects on daphnids or chironomids after 14 and 30

day exposure periods, and no effects of chronic exposure at 10 or 1,000 Ilg/L on daphnid

adult or instar survival. For the 7-day chronic toxicity test, the treatment levels of 0, 10,

and 1,000 ).lg/L atrazine had: 90% gravid females at all treatment levels; the number of

adults >1,500).lm long were 37.5 ± 5.4,46.7 ± 9.0, and 45.3 ± 6.0, respectively; and the

number of instars <1,500).lm long were 28.7 ± 13.5,20.5 ± 1.7, and 20.0 ± 5.3,

respectively; there were no significant differences at pSO.05. (Numbers are means ±

standard deviations with four replicates.)

Phytoplankton lethality tests found that atrazine exposure of 1,000 ).lg/L resulted in

significant (psO.05) biomass reduction. Dry weight algal biomasses, with acute exposure

after 30 days in the microcosm, were (mean ± standard deviation with four replicates):

5.8 ± 0.8 mg at 0 Jlg/L

6.9 ± 0.5 mg at 10 Ilg/L

6.6 ± 0.9 mg at 100 Jlg/L

5
3.7 ± 0.5 mg at 1,000 flg/L

Gross primary productivity (GPP) gradually increased over time in most microcosms

(Figure 1). There was some initially wide variation within treatments which the author

attributes to differences in macrophyte establishment. Immediately following atrazine

treatment, there was a decline in GPP, and dissolved oxygen and community respiration

decreased as well. Reductions in GPP relative to controls were 23, 32, and 91% in the

10, 100, and 1,000 flg/L treatments, respectively. The 10 and 100 flg/L treatments

recovered at 7 days and remained similar to controls until the experiment ended, but the

1,000 flg/L treatment never recovered.

Macrophyte biomass was not affected by 30 days of atrazine exposure at 10 or 100 flg/L,

but there were significant (p:S;0.05) reductions of about 50% in the 1,000 flg/L treatments.

Dry weight total biomass results were (mean ± standard deviation with four replicates):

1.64 ± 0.56 g at 0 flg/L

1.94±0.71 gat 10 flg/L

1.45 0.20 gat 100 flg/L

0.86 ± 0.24 g at 1,000 flg/L

Water quality measurements showed some response to atrazine treatment. Conductivity

in the 1,000 flg/L treatments was increased about 20% over controls and pretreatment

values (Figure 2); other treatment levels showed no significant (p:S;0.05) differences from

controls. Additionally, pH was lower in the 1,000 flg/L treatment microcosms, and on 3

ofthe 4 posttreatment weeks was significantly (p:S;0.05) different from control (Figure 3).

Again, other treatment levels showed no significant differences. Alkalinity

measurements at 30 days were doubled in the 1,000 flg/L treatments compared to the

controls, a significant (p:S;0.05) difference, but the other treatments did not have

significant differences. Final (30 day) alkalinity measurements (in mg as CaC03) were

(mean ± standard deviation with four replicates):

104.0 ± 27 at 0 flg/L

96.0 ± 46 at 10 flg/L

142.7 ± 65 at 100 flg/L

229.0 ± 39 at 1,000 flg/L

The author describes a possible mechanism for the microcosm feedbacks resulting in

these water quality changes. Total nitrogen (TN) generally increased over time in all

microcosms, by about 28% over the 30 days ofthe experiments, with no noticeable

effects due to atrazine treatment. On the other hand, total phosphorus (TP) tended to

decrease about 50% over that time period, in controls and at 10 and 100 flg/L atrazine

treatment levels. At 1,000 flg/L atrazine, TP remained simi lar to pretreatment levels.

Total organic carbon in the microcosm sediments showed no significant (p:S;0.05)

6
differences between treatments and controls at the end ofthe 30 day experiments, with a

range in dry weight of about 8-12%.

Microbial activity measures were not affected significantly (p:S0.05) by atrazine

exposure. Only 1,000 /lg/L tests were performed. Uptake rates ofINT-F (2-(Piodophenyl)-3-(p-phenyl-5-phenyl) tetrazolium-formazan, for determining electronic

transfer system activity), PNP (p-nitrophenol, for determining monophosphatase

activity), O2, and glucose were measured per gram dry weight of hydrosoil (except for

glucose uptake rates, which are expressed per gram wet weight hydrosoil). Results were

(mean ± standard deviation with four replicates):

/lg INT-F/g/hr: 2,616 ± 87 (treatment); 2,615 ± 162 (control)

/lg PNP/g/hr: 517 ± 28 (treatment); 481 ± 38 (control)

IlL Oz/g/hr: 90 ± 13 (treatment); 89 ± 9 (control)

/lg glucose/g/hr: 2.6 ± 0.3 (treatment); 2.7 ± 0.4 (control)

7
LJ L.L-.l

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\ ., ._--~._------ ..•.---. c

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' \.:-_.-._..-~_... -_ ..-.......... Q ..L1..Li.JJLL1 LL: LLL I LJ IL

20 r-r-n-rrrrnTrnTT1'nTTITl TT"'rrrl

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Figure 1. Influence of atrazine on gross primary productivity. All values are means of four

replicates. Asterisks denote significant differences (p~0.05) from control; mean differences

determined by ANOVA, based on a nested design. Nominal atrazine concentrations were control

(open circle, 0), 10 IJ.g/l, (solid square, _),100 prg/l, (open square, D), and 1,000 prg/L (solid circle, e).

Day 0 denotes the day of pesticide treatment.

8
60;)

t

;;50

.§

'~ SOO t

'- E

,= 4~)

;;

8 400

L.

350 L.-

!

3('l()

o

Figure 2. lnfluence of atrazine on conductivity. All values are means of four replicates. Asterisks

denote significant differences (p~0.05) from control; mean differences determined by ANOVA, based

on a nested design. Nominal atrazine concentrations were control (open circle, 0), 10 f1g/L (solid

square, _),100 f1g/L (open square, 0), and 1,000 f1g/L (solid circle, .). Day°denotes the day of

pesticide treatment.

9

-------------------
1(;

1

t-

'!'

9 ,;

-j

.L ...

o

,/

./

•

.~..L ... __I . ~1

l

I

Figure 3. Influence of atrazine on pH. All values are means of four replicates. Asterisks denote

significant differences (p:::S0.05) from control; mean differences determined by ANOVA, based on a

nested design. Nominal atrazine concentrations were control (open circle, 0),10 Itg/L (solid square,

_),100 /lglL (open square, 0), and 1,000 Itg/L (solid circle, .). Day 0 denotes the day of pesticide

treatment.

10
From study

Table 5.

Data Summary Table: P-values are shown for significant difference from controls.

Effect Magnitude of Nominal Day ofStudy Time Did Comments

Effect relative Atrazine Effect Weighted Recovery

to controls Concentration Occurred Average Occur - if

(pg/L) Atrazine so, when

Concentration (see

! from Day 0 to comment)?

I When Effect j v I _--I ..__. f-- .._._--t--:l=.-...:..... ',A:.:::.c..:."...:"'.::.:.-u + --.-----I--~--.-

i:~rea:e in- ~~~~~6~~t-----+-1.0--f.l.-g.I-L-_---l--N-A-_-~.-j---.----i Effect was

macrophyte No effect 100 NA evaluated at

biomass ( >0.05) end ofstudy.

I 50% (p<0.05 1.000 ug/L Dav 30 NA

Decrease L..~~~<0.05 .l.QJl:.g/L Day 2 NA NA , From Fig.l

;~~~sar 32% (p>0.05 lQQ..gg/L II Day 2 I ~~ ~IN~A I and text p.

Yt"t 91% (.p:S0.05) 1,000 IIgiL Day") , No 480. roduc IVIy' I'"

Change In f-N'o effect II NA- kA - - NA-----+----------J

10 IIgll From study daphnid 1 (p>0.05) I'" j

I f-.- -----+'-----.- Table 3 and

reproduction No effect I 1000- ,Z INA NA ~NA text p. 479.

* (p>0.05) .---~g --=t ----+--------1

No effect 10 IL NA NA NA

Change in (p>0.05) IJ,g ---~--- -._~--=E- ----- From study

sediment No effect ,I NA

~~~:7~~1 ~~~;;t II :OoOo:g/L/L=+~~----+NA --- ~~~----l ~:~1;.74~~~ __~...:..(P.....:>_O~_ '_~!_ _'~_ ' ---'

No effe~t l NA

(p>0.05) 10 1!.. g/L : NA --nNA _ ..... __I NA From Fig.2 Increase in . - r--- .. . No effect I NA and text p. conductivitv 100 IIglL NA I NA ') I (p>0.05) i I'" I L- ---1 481.

20% (p:SO.02..L+ 1,000 Ilg/L I Day 7 I No ---j--__ ~o oeffioe5,~t I 10 Jlg/L ~A NA j NA M' d

\\.'> . ') _~____ ~_ I ! agmtu es Decrease in ----t

pH I ~o effect I 100 IL : NA I NA INA I estimated

>_0.05t_~__!g -----J------- . 1 from Fig.3.

_____ ..10% (p<0.05) i 1,000 J.lglL . Da. 30 NA -+NQ...___ 1 _

No effect 1 10 giL I NA NA I NA (p>O.O~ J.l . . ......-+-1 _

. No effect NA

,Ikalin;ty 1:0% ~~ ~-·---1· NA - - EffeotW",!' Table 6.

( <0.05) I 1,000 J.lg/L Day 30 I measured at

p- I . _-L..--. I I end ofstudy I ~----_'-:

Note: NRc=Not Reported, NA=Not Applicable

a Atrazine concentrations were not measured; therefore, time-weighted average values were not

detelmined.

11
No atrazine concentration measurements were taken in the microcosms, so the actual

concentrations are unknown. Time weighted averages were assumed equal to nominal

due to the lack of information.

Reviewer Comments:

This study provided good replication, with four replicates at each ofthree treatment

levels plus four controls. A wide range of measurements were taken on various

components ofthe microcosm systems. However, the study is strongly limited by lack of

atrazine concentration data. The atrazine was added as part of a soil slurry, which was

not mixed into the microcosms but allowed to naturally settle from the top, in an attempt

to recreate natural runoff conditions. The methodology is also likely to result in atrazine

concentrations much different than nominal, probably smaller concentrations due to

adsorption to the soil particles in the slurry. This may be why so little effect was found

from the 10 and particularly the 100 Ilg/L atrazine treatments.

As with any microcosm study, there are limitations to how realistic the laboratory

conditions could be in recreating a natural aquatic environment. The microcosms in this

study contained a reasonable set of aquatic ecosystem components, including

macrophytes, but still may not be complex enough to stand in for a real pond. One ofthe

primary shortcomings is the ill-described, apparently somewhat short acclimatization

process. It does not appear that the microcosms had reached steady state when the

experiment commenced; for example, the author notes that some within-treatment

variability in OPP was probably due to differences in macrophyte establishment.

Description of Use in Document (QUAL, QUAN, INV): QUAL

Rationale for Use:

Study evaluated relevant endpoints and contributes to the weight of evidence regarding

potential effects of Atrazine on aquatic plants. However, atrazine concentrations were

not measured.

Limitations of Study: No atrazine concentrations were measured, but are probably far

from nominal due to surface-only application within a soil slurry. Microcosms may be

too simplified to apply to natural environments, and may not have reached steady state by

the start ofthe experiment.

Primary Reviewer: Sara Hendrix, M.S., Staff Scientist, Cambridge Environmental Inc

Secondary Reviewer: Brian Anderson

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