### Open Literature Review Summary

ChemicalName: Atrazine 

### CAS No: 1912-24-9



### ECOTOX Record Number and Citation: Downing, 

### H. F., M. E. DeLorenzo, M. 

### H. Fulton, G. 1. Scott, C. J. Madden, and J. R. Kucklick. 2004. Effects ofthe agricultural pesticides atrazine, chlorothalonil, and endosulfan on south Floridamicrobial assemblages. Ecotoxicology. 13: 245-260. (MRID 48261110)


### Purpose ofReview: Determine potential effects ofatrazine on an aquatic community. Endpoints from this study contributeto the weight-of-evidence regarding effects ofatrazine on aquatic plantcommunities and associated determination ofa Level ofConcern (LOC).



### Date ofReview: October 7,2009


### Summary ofStudy Findings
Purpose ofStudy: This study examines the potential effects ofatrazine onphytoplankton and zooplankton collected from two locations along a southernFloridacanal. Chlorothaloni1 and endosulfan effects onthe aquatic microbial assemblages were investigated separately; those results are notreported here.



### Methods: Microbial assemblages (algae, bacteria and protozoa) were collected from two locations (Sites C and E) along the C-111 canal, located in Miami-Dade County, Florida, during October (wet season) 1999 and February (dry season) 2000 

### (Figure 1). Site C was surrounded on all sides by agricultural fields used primarily for tomato production, while Site E,**~5** miles downstream from Site C, was surrounded mostly by sawgrass marsh. Artificial substrates [polyurethane foam units (PFUs), each 4 x 5 x 6 cm3], attached to a 2-m nylon rope via cable-ties, were positioned at a depth of30 cmbelow the water surface, forty-eight substrates per collection site. Microorganisms were allowed to colonize the artificial substrates for 7 days (168 hours). The substrates were then collected, beneaththe water surface, into I-gallon, plasticjars; eight substrates perjar. Thejars, containing substrates plus canal water, were sealed and transported on ice to the NOAAlNational Ocean Service (NOS) Charleston (South Carolina) Laboratory. Priorto transport, microorganisms were "extracted" from some ofthe artificial substrates (PFUs, numbernot specified) on site immediately aftercollection by gently squeezing the PFU contents - 1 -



# ' ••••••111 2087821**(~60** 

### mL) into individual, sterile samplejars, with additional PFUs "extracted" following transportto the laboratory. Canal water (24 L) was collected from each site into stainless steel cans (size not specified) for use during atrazine exposure experiments. Additional canal surface water (volume not specified) was collected from each site into l8-L stainless steel containers at five unspecified intervals during the 7-day colonizationperiod for atrazine analysis as described below.

Atrazine exposure. Eachtest system consisted ofa 600-mL glass beaker containing one colonized artificial substrate (PFU) and 500 mL offiltered (0.22-llm, 

### Gelman Maxi Capsule Filter) canal water (respective to PFU collection site) amended with nutrients according to ASTM (1996) standard protocols for freshwater mediumpreparation. Atrazine (purity 96%), in acetone, was applied at 20 and 200 Ilg/L; final co-solvent concentration 0.01 %. Control systems treated with acetone at 0.01% were also prepared. All treatments (0, 20 and 200 Ilg/L atrazine) were replicated intriplicate. Following treatment, the beaker contents were stirredto mix. Incubation conditions were notreported. 15 mL water samples were collected for atrazine analysis randomly from the beakers at 0, 24, and 168 hours posttreatment. Artificial substrates (PFUs) were removed at 24 and 168 hours posttreatment and the microorganisms collected (extracted) as described above. Extractedmicroorganisms were homogenized by gently stirring, then aliquots were removed and analyzed for chlorophyll 

### a content, photosynthetic activity (carbon assimilation), bacterial abundance, heterotrophic bacterial productivity (leucine incorporation) and species composition as described below.

Atrazine in canal surface water. 2-3 Lwater samples were filtered (0.7-llm, 

### 90-mm glass fiber, Whatman type GF/F) underpressure, loaded onto pre-conditioned (methylene chloride, acetone) 0.5-g Isolute ENV+ solidphase extraction (SPE) cartridges, and stored frozen (temperature, interval notreported) until elution. Atrazineresidues were eluted with acetone, followed by methylene chloride. Extracts were evaporated undernitrogen, thenresulting residues were reconstituted inhexane and, as needed, cleanedup viaflorisil micro-column. Samples were again evaporatedundernitrogen, with resulting residues reconstituted in isooctane and analyzed using GC-MS with electron ionization (EI) underthe following conditions: J&W Scientific DB5ms GC column (0.25 mm x 60 m, 0.25 11m 

### film thickness), injection volume 2.0 ilL, flow rate 1.5 mL/min., velocity 32 cm/sec., collision gas methane, temperature program of60°C for 1 min. : ramp to 150°C at 25°C/min. : ramp to 240°C at 5°C/min. : ramp to 300°C at 15°C/min.: hold for 3 min., total run time 29.60 min., monitoring m/z 200 and 215, with atrazine-d4as an internal standard (m/z 205); limitofdetection was not reported.

Verification ofatrazine treatmentrates for exposure tests. The 0-, 24- and 168-hourtest system water samples (15 mL) were separately loaded onto pre-conditioned (methanol, deionized water) l-g octadecyl (C18) SPE cartridges (Burdick and Jackson). After sample loading, the cartridge was rinsed with water, dried undervacuum for 5 minutes and atrazine was eluted with ethyl acetate. The resulting eluate was dried over anhydrous sodium sulfate (Na2S04), evaporated undernitrogen, with the residues reconstituted in isooctane and analyzed by GC/MS-EI as described above. - 2 -Chlorophyll 

### a. Microorganism samples (10 mL) were filtered (GF/F), thenthe filters were placed in vials with saturated magnesium carbonate (MgC03) solution (1 mL) and stored frozen (-20°C, interval not specified) until analysis. Priorto analysis, samples were extracted with acetone (final concentration 90%), shaken, refrigerated overnight, shaken and again refrigerated overnight (total extractioninterval 48 hours), then broughtto room temperature in darkness and analyzed using a fluorometer (Sequoia-TurnerModel 450).
Phototrophic carbon assimilation (primary productivity). Microorganism samples (5 mL) were treated with [14C]sodiumbicarbonate (NaH14C03, concentrationnotreported), then maintained at 

### in situ light and temperature regimes (not further detailed) in an incubator for 24 hours. Following incubation, the samples were filtered**(0.45-~m,** cellulose, nitrate), then the filters were rinsed with filtered**(O.2-~m)** canal water (respective to microorganism collection site), placed in scintillation vials with 10% HCI and allowed to sit in darkness overnight. Thereafter, scintillation fluid was added and the samples were equilibrated overnight and analyzed for total radioactivity using LSC.
Phytoplankton (microalgae) community composition. Changes in community based onphotopigment composition were investigatedusing October 1999 microorganisms exposedto atrazine (20 and 200**~g/L)** for 24 hours. Microorganism samples (10-20 mL) were filtered (25-mm, GF/F) and the filters stored frozen (-80°C) until extraction. Samples were extracted with icecold acetone and sonication (1 minute), withresulting extracts stored frozen (-20°C) overnight. The samples were centrifuged (2,795 x g, 10 minutes, 4°C), then an aliquot (1 mL) was filtered**(0.2-~m)** into a nitrogen-filled, amber 1.8-mL HPLC vial and stored frozen (-80°C) until analysis by reverse-phase HPLC underthe following conditions: VYDAC 201TP C18 column (4.6 x 250 mm, 5**~m** particle size, 300

### Apore size), methanolic mobile phases (Van Heukelem 

### etal., 1992), inline diode array detector (method not further described). Pigments were identified based on comparisons ofabsorbance spectra and againstreference standards.
Community composition was determined viaidentification ofphototrophic taxain gluteraldehyde preserved samples (see bacterial abundance below) using an Olympus BX-50 epifluorescence microscope with magnification at 500.

Bacterial abundance was determined via acridine orange direct counts; method ofHobbie 

### etal. (1977) as modified by Kepner and Pratt (1994). Microorganism samples (5 mL) were preserved with gluteraldehyde (final concentration 2%), then stored in darkness at4°C until analysis. Preserved samples were diluted with filtered**(0.2-~m)** deionized water, then stained, in darkness, with acridine orange (100**~g/L,** final concentration) for 3 minutes. The stained samples were filtered**(0.2-~m** Poretics PCTE blackpolycarbonate membrane), then bacterial counts were conducted using an Olympus BX-50 epifluorescence microscope, with magnification at 1,250, and equipped with a superhigh-pressure mercury lamp underblue light excitation (470-490 nm).

- 3 -Leucine incorporation (heterotrophic bacterial productivity). Microorganism samples (5 mL) were treated with eH]leucine (105 nmol/L, final concentration) and incubated in darkness at in situ temperature (not further detailed) for 60 minutes. Addition of50% trichloroacetic acid (TCA) stopped leucine incorporation, and the samples were extracted in ahot waterbath at 80°C for 15 minutes. Extracted samples were cooled, then filtered (0.45-llm cellulose nitrate) under low vacuum. Filters were rinsed with ice-cold 5% TCA, followed by ice-cold 80% ethanol, then placed in scintillation vials, dissolved with ethyl acetate and analyzed for total radioactivity using LSC.

Statistical analyses. Response values were analyzed for variance heterogeneity and normality (Shapiro WilkW test, JMP software) and, ifrequired, results were transformedto meet assumptions ofparametric tests. A multi-factor, multivariate analysis ofvariance (MANOVA) was usedto detect differences in microbial assemblages among seasons, collection sites, treatments and length ofexposure. Average number oftaxaperPFU were analyzed using twoway ANOVA. Forboth MANOVA and ANOVA analyses, apriori orthogonal contrasts were used to detect significant differences between treatments and controls. Photo-pigmentresults (normalizedto chlorophyll a) and changes in relative abundance ofdominanttaxa groups were analyzedusing two-factor MANOVA, with collection site and treatment as main effects. Genera were grouped as follows: (1) Bacillariophyta, (2) Cryptophyta, Chrysophyta and Dinophyta, (3) Cyanophyta, (4) Heterotrophic protists, and (5) Chlorophyta and Euglenophyta.

Results: In the canal water at sites C (agricultural site) and E (sawgrass marsh site), atrazine was detected at 18.92 ± 1.20 ng/L and 24.5 ± 3.77 ng/L, respectively, in October 1999 (wet season), and at 32.68 ± 1.24 ng/L and 11.3 ± 1.01 ng/L, respectively, in February 2000 (dry season) indicating the microbial assemblages were pre-exposed to atrazine during the 7-day colonization period 

### (Table 1).
During the acute exposure experiments, measured atrazine concentrations at the 20 Ilg/L application rate were 119-124% ofnominal at time 0, decreasing to 82-101% ofnominal at 24 hours and 69-86% ofnominal at 168 hours 

### (Table 2). At the 200 Ilg/L application rate, atrazine was detected at 56-92% ofnominal at time 0, 59-69% ofnominal at 24 hours and 41-48% of nominal at 168 hours. Any analysis ofwater from the untreated control test systems was not reported.

At the 200 Ilg/L treatment rate, atrazine significantly (p<0.05) reduced phytoplankton biomass**(~30-40%,** as chlorophyll a) and primary productivity**(~60%,** as carbon assimilation). With the exception ofthe February 2000 microorganisms, where biomass was decreased only**~5%** after 24 hours (Figures 2-3), atrazine exposure reduced biomass and primary productivity regardless

- 4 -

### ofsite (C or E), collection season for microbial assemblages (Oct. or Feb.), or exposure duration (24 or 168 hours).

At the 20**~g/L** 

### treatment rate, phytoplankton biomass and primary productivity were not significantly decreased**(:S~** 

### 10%,**p~0.05),** 

### when responses were pooled for site, season and length of exposure (Figure 2). However, in a comparison of seasonal variability, phytoplankton biomass collected in Oct. was significantly (p<0.05) decreased**~35%,** 

### as compared to controls, after 168 hours ofexposure, while biomass for phytoplankton collected in Feb. was stimulated**~+20%** 

### (not significantly different from control) after 168 hours (Figure 

### 3).
Two-way MANOVA determined significant (p<O.OOOI) differences in microbial assemblages between the two collection sites, withthe artificial substrates colonized at site E (sawgrass marsh site) having more taxa (n = 

### 54), as compared to substrates colonized at site C (agricultural site, n

= 

### 43, p. 254). Relative abundance of green algae (chlorophytes + euglenophytes) and cyanobacteriataxawere higher at site C (30% and 23% oftaxa, respectively), than at site E (26% and 15%, respectively). While diatoms and heterotroph taxa were higher at site E (33% and 17%, respectively), than at site C (28% and 12%, respectively).

Atrazine exposure significantly (p<0.05) decreased total taxa (phytoplankton + heterotrophs, sites combined) 10% and**~25%,** 

### as compared to controls, after 24 hours at both the 20 and 200**~g/L** 

### treatment rates, respectively (Figure 

### 4). However, the extent of the effects varied according to collection site with total taxa decreased 14% and 36% for microorganisms from sites C and E, respectively, at the 200**~g/L** 

### treatment rate; site results for the 20**~g/L** 

### rate were not presented. Based on the reviewer's interpretation of Figure 

### 4, diatoms, bluegreen algae, green algae and brown algae were decreased**~20%, ~25%,** **~30%** 

### and 75%, respectively, at the 200**~g/L** 

### rate, as compared to controls, while diatoms, green algae and brown algae were decreased**~15%, ~1O%**  

### and**~25%,** 

### respectively, at the 20**~g/L** 

### rate with no effect to bluegreen algae, and no effect to heterotrophs at either treatment rate. The study authors reported that green algae and chrysophytes decreased 4% and 5%, respectively, at the 200**~g/L** 

### rate, while diatoms and heterotrophs each increased by 4%; however, that statement, specifically in regards to diatoms, appears contradictory to the results depicted in Figure 4.


### Pigment analysis found significant (p<0.0001, two-way MANOVA) differences in algal pigment composition between the two collection sites, with fucoxanthin the most abundant pigment at both sites; quantitative results were not presented. Concentrations (normalized to chlorophyll a) 

### ofthe pigments fucoxanthin, chlorophyll eland c2 were reported to be significantly higher (p value not reported) at site E, while chlorophyll b 

### was more abundant at site C. Concentrations of pigments violaxanthin, lutein and canthazanthin were not reportedly significantly (p value not reported) different between the two collection sites. Atrazine, at 200**~g/L,** 

### decreased levels of fucoxanthin**(~20%)** 

### and increased chlorophyll b**(~26%)** 

### in phytoplankton from site C, while resulting in almost no change «10%) in pigment levels in phytoplankton from site E; however, responses were reportedly not significantly different from controls (p value not reported).
- 5 -

### Bacterial abundance was significantly decreased**(~20%,** 

### p<0.05) at the 200 f.lg/L treatment rate, when responses were pooled for site, season and length ofexposure, butthere was no observable decrease atthe 20 f.lg/L rate (Figure 2). Bacterial productivity (as eH]leucine incorporation) was stimulated at both the 20 f.lg/L**(~1** 

### 0-40%, not statistically significant, p2:0.05) and 200 f.lg/L**(~40-100%,** 

### statistically significant, p<0.05) treatmentrates (Figures 2 and 

### 5).

The study authors proposed that chronic exposure to "background" levels ofatrazine in the canal may have selected for species that were more resistant, resulting inthe reduced responses seen in these experiments at the lower 20 f.lg/L atrazine treatmentrate.



### - 6 -

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### "*,, = significantly (p<0.05) different from control. Low and High treatment rates were 2 and 20 I1glL, respectively, for chlorothalonil, and 1 and 10 I1glL, respectively, for endosulfan. Figure 2 from Downing etal. 2004.



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Figure 3. Mean responses (pooled for site) ofchlorophyll a (/!glL) to atrazine at 20 /!g/L ("Low") and 200 /!glL ("High") at each incubation interval, with standard deviation (vertical bars); 

### "*,, = significantly (p<0.05) different from control. Top = 24 hr posttreatment, bottom = 168 hr. Low and High treatment rates were 2 and 20 /!glL, respectively, for chlorothalonil, and 1 and 10 /!glL, respectively, for endosulfan. Figure 4 from Downing etal. 2004.

- 9

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Figure 4. Average number oftaxa found in each atrazine treatment (20 or200 f.lg/L) after 24 hours of exposure with microorganisms collected in October 1999 at both sites; 

### "*" = significantly (p<0.05) different from control. Low and High treatment rates were 2 and 20 f.lg/L, respectively, for chlorothalonil, and 1 and 10 f.lg/L, respectively, for endosulfan. Figure 4 from Downing etal. 2004.

- 10

-Chh,rotlu;",,,il Lnamulfan o loll.' cl:r.:c-ntr.ui-I:f"'J Ii hi,£h o;:t;ln"nLr.u.ktn**:.~h(l~** 2j b**:.l.~·.,O~** 

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## " .=**.~** 1.1:'1-.,(15 II**.~** .U4·:,(14 E!**(I.[~·.f(.n** C'>!llJ'>:o1 Figure 5. Mean responses (pooled for site and season) ofbacterial incorporation of[3H]leucine (CPM) to atrazine at 20 p.g!L ("Low") and 200 p.glL ("High") at each incubation interval, with standard deviation (vertical bars); ); 

### "*,, = significantly (p<0.05) different from control. Top = 24 hr posttreatment, bottom = 168 hr. Low and High treatment rates were 2 and 20 p.g!L, respectively, for chlorothalonil, and 1 and 10 p.glL, respectively, for endosulfan. Figure 3 from Downing etal. 2004.

- 11

-Table 1. Atrazine concentrations (ngIL, mean ± st. dev., n = 5) in canal surface water during I-week microorganism colonization period (from Table I ofstudy). Site C l Site El Season2 Mean ± St. Dev. (Range) Mean ± St. Dev. (Range) Oct-99 (wet season) 18.92 ± 1.20 (17.2-20.0) 24.5 ± 3.77 (18.8-28.4) Feb-OO (dry season) 32.68 ± 1.24 (31.2-33.9) 11.3 ± 1.01 (10.2-12.8) 1 Site C was surrounded on all sides by agricultural fields used primarily for tomato production, while Site E,**~5**

miles downstream from Site C, was surroundedmostly by sawgrass marsh. 2 Florida agricultural pesticide application heaviest during dry season (December-April) and lightest during wet

season (May-November).

Table 2. Measured atrazine concentrations (flgIL, mean ± st. dev., n not specified) for treated microorganism exposure experiments (from Table 4 ofstudy).

200 112 ± 25.2 117 ± 2.54 +4.0 81.3 ± 0.25 28.0 Season

Oct-99

Feb-OO Nominal (Jig/L)

20 200 20 Ohr
23.8 ± 6.14 184 ± 3.37 24.8 ± 3.80 24 hr
16.4 ± 0.12 137 ± 0.03 20.2 ± 0.27 % Decreasel

31.3 26.0 18.3 168 hr
13.7 ± 0.06 95.3 ± 2.29 17.1±0.49 % Decrease l

42.6 48.4 31.0

1 Decrease in average atrazine concentration from that detected at 0 hr, except for Feb-OO 200 Jig/L treated at 24

hours which was an increase from 0 hr.



### - 12

-

### Data Summary Table: Natural microbialassemblages incubatedindoors (in open beakers) Time Weighted Magnitude Nominal Day of Study AverageAtrazine Did Effect of Effect Atrazine Effect Concentration Recovery Comments relative to Concentration from Day Oto Occur-if controls (p.g/L) Occurred When Effect so, when? Occurred Magnitude ofeffect 20 J.lg/L; Not applicable; estimated from Phytoplankton No observed measured 23.8- pooled mean of Not applicable Not Figure 2. Effectnot biomass effect 24.8 J.lg/L 24- and 168- applicable statistically (chlorophyll a, hourresponses significant J.lg/L); pooled (p2:0.05).

mean response Magnitude ofeffect for site, season Not applicable; mean 111.9 J.lg/L1; estimated from and length of**~30%** 200 J.lg/L; pooled mean of Experiment Figure 2. exposure decrease measured 112- 24- and 168- Oct. 122.5 J.lg/L, terminated at Effect statistically 184J.lglL hourresponses Feb. 101.3 J.lg/L 168 hours. significant (p<0.05). Magnitude ofeffect Primary 20 J.lg/L; Not applicable; estimated from productivity No observed measured 23.8- pooled mean of Not applicable Not Figure 2. Effect not (carbon effect 24.8 J.lg/L 24- and 168- applicable statistically assimilation, hourresponses significant mg elL/day); (p2:0.05). pooled mean Magnitude ofeffect response for Not applicable; mean 111.9 J.lg/L; Experiment estimatedfrom site, season and**~60%** 200 J.lg/L; pooledmean of Figure 2. length of decrease measured 112- 24- and 168- Oct. 122.5 J.lg/L, terminated at Effect statistically exposure 184 J.lg/L hourresponses Feb. 101.3 J.lg/L 168 hours. significant (p<0.05).

20 J.lg/L; No (effect**~35%** observed at Magnitude of Phytoplankton decrease measured 23.8 168 hours 15.8 J.lg/L end of effects estimated biomass J.lg/L experiment) from Figure 3. (chlorophyll a, Effects statistically J.lg/L); pooled 200 J.lg/L; significant for site (Oct99)**~30%** measured 184 24 hours 160.5 J.lg/L No (p<0.05). decrease J.lg/L Magnitude ofeffect 20 J.lg/L; estimated from Not Figure 3. Effectnot Phytoplankton No effect measured 24.8 Not applicable Not applicable applicable statistically biomass J.lg/L significant (chlorophyll a, (p2:0.05). J.lg/L); pooled No (effect for site (Feb 00) 200 J.lg/L; Magnitude ofeffect**~40%** measured 112 168 hours 101.3 J.lg/L observed at estimated from decrease end of Figure 3. Effect J.lg/L experiment). statistically

- 13

-Time Weighted Magnitude Nominal Day of Study AverageAtrazine Did Effect olEffect Atrazine Effect Concentration Recovery Comments relative to Concentration from Day Oto Occur-if controls (pg/L) Occurred When Effect so, when? Occurred significant (p<0.05).**~15%** 2Ollg/L; Only interval Bacillariophytes decrease measured 23.8 24 hours 20.1 Ilg/L analyzed. Magnitude of (diatoms, Ilg/L effects estimated average 2OOllg/L; from Figure 4; number)2**~20%** measured 184 24 hours 160.5 Ilg/L Only interval SSNR3

. decrease4 Ilg/L analyzed. 2Ollg/L; Not Cyanophytes No effect measured 23.8 Not applicable Not applicable applicable Magnitude of (b1uegreens, Ilg/L effects estimated average 2OOllg/L; from Figure 4; number)2**~25%** measured 184 24 hours 160.5Ilg/L Only interval SSNR. decrease Ilg/L analyzed.**~5%** 2Ollg/L; Only interval Chlorophytes + decrease measured 23.8 24 hours 20.1 IlglL analyzed. Magnitude of Euglenophytes Ilg/L effects estimated (greens, average 2OOllg/L; from Figure 4; number)2**~30%** measured 184 24 hours 160.5 Ilg/L Only interval SSNR. decrease4 Ilg/L analyzed.
Chrysophytes +**~25%** 2Ollg/L; Only interval Cryptophytes + decrease measured 23.8 24 hours 20.1 Ilg/L analyzed. Magnitude of Dinophytes j..lg/L effects estimated (browns, 2OOllg/L; from Figure 4; average**~75%** measured 184 24 hours 160.5 j..lg/L Only interval SSNR. number)2 decrease4 j..lg/L analyzed.**~10%** 20 j..lg/L; Only interval Total decrease measured 23.8 24 hours 20.1 Ilg/L analyzed. Magnitude of phytoplankton j..lg/L effects estimated (average 200 j..lg/L; from Figure 4; number)2**~30%** measured 184 24 hours 160.5 j..lg/L Only interval SSNR. decrease j..lg/L analyzed.
20 j..lg/L; Only interval No effect measured 23.8 Not applicable Not applicable analyzed. Magnitude of Heterotrophs Ilg/L effects estimated (average from Figure 4; number)2 200 j..lg/L; Only interval SSNR. No effect4 measured 184 Not applicable Not applicable analyzed. Ilg/L Total**~10%** 20 j..lg/L; 24 hours 20.1 Ilg/L Only interval Magnitude of

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-Time Weighted Magnitude Nominal Day of Study AverageAtrazine Did Effect of Effect Atrazine Effect Concentration Recovery Comments relative to Concentration from Day Oto Occur-if controls (pglL) Occurred When Effect so, when? Occurred microorganisms decrease measured 23.8 analyzed. effects estimated (average Jlg/L from Figure 4. number)2 200 JlglL; Effects statistically**~25%** Only interval significant decrease5 measured 184 24 hours 160.5 JlglL analyzed. (p<0.05). JlglL Magnitude ofeffect 20 JlglL; Not applicable; estimated from Bacterial No observed pooledmean of Not Figure 2. Effectnot biomass effect measured 23.8- 24- and 168- Not applicable applicable statistically (abundance, 24.8 Jlg/L hourresponses significant cells/mL); (p2:0.05). pooledmean response for Magnitude ofeffect site, season and 200 JlglL; Not applicable; mean 111.9 IlglL; Experiment estimated from length of**~20%** measured 112- pooledmean of Oct. 122.5 Jlg!L, terminated at Figure 2. exposure decrease 184 Jlg/L 24- and 168- Feb. 101.3 Jlg!L 168 hours. Effect statistically hourresponses significant (p<0.05). Magnitude ofeffect Bacterial 20 Jlg!L; Not applicable; mean 17.5 JlglL; Experiment estimated from productivity No effect measured 23.8- pooledmean of Oct. 15.8 Jlg!L, terminated at Figure 2. Effectnot 24- and 168- statistically (leucine 24.8 Jlg/L hourresponses Feb. 19.2 Jlg/L 168 hours. significant incorporation, (p2:0.05). cpm); pooled mean response Magnitude ofeffect for site, season 200 JlglL; Not applicable; mean 111.9 1lg!L; Experiment from article text (p. and length of**~46%** measured 112- pooledmean of Oct. 122.5 Jlg!L, terminated at 253). exposure stimulation 184 Jlg/L 24- and 168- Feb. 101.3 JlglL 168 hours. Effect statistically hourresponses significant (p<0.05, Figure 2). Magnitude ofeffect 20 JlglL; Experiment estimated from Bacterial No effect measured 23.8- Not applicable Not applicable terminated at Figure 5. Effectnot productivity 24.8 JlglL 168 hours. statistically (leucine significant incorporation, (p2:0.05). cpm); pooled Magnitude ofeffect mean response for site and 2OOIlg!L; mean 111.9 IlglL; Experiment estimated from**~100%** Figure 5. season stimulation measured 112- 168 hours Oct. 122.5 Jlg/L, terminated at Effect statistically 184 Jlg/L Feb. 101.3 JlglL 168 hours. significant (p<0.05). Mean ofTWAs for October 1999 and February 2000 expenment (attached Worksheet).

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-2 Average numbers estimated from Figure 4. For Control: bacillariophytes (diatoms) = 15, cyanophytes (bluegreens) = 8, chlorophytes + euglenophytes (greens) = 13, chrysophytes + cryptophytes + dinophytes (browns) = 4 [yielding total phytoplankton = 40], heterotrophs = 7 [yielding total microorganisms = 47]. For "Low" atrazine treatment: bacillariophytes = 13, cyanophytes = 8, chlorophytes + euglenophytes = 12, chrysophytes + cryptophytes + dinophytes = 3 [yielding total phytoplankton = 36], heterotrophs = 7 [yielding total microorganisms = 43]. For "High" atrazine treatment: bacillariophytes = 12, cyanophytes = 6, chlorophytes + euglenophytes = 9, chrysophytes + cryptophytes + dinophytes = 1 [yielding total phytoplankton = 28], heterotrophs = 7 [yielding total microorganisms = 35]. 3 Statistical Significance, as comparedto controls, was Not Reported. 4 The study authors reportedthatrelative abundance ofchlorophytes + euglenophytes (greens) and chrysopytes

decreased4% and 5%, respectively, while diatoms and heterotrophs each increasedby**~4%** (p. 254); the reviewer did not agree withthis conclusion, based on the data shown in Figure 4. 5 Total microorganisms from sites C and E were reduced 14% and 36%, respectively, as compared to controls (p.

254).

- 16 -Reviewer Comments: 

### No analysis for atrazine or other pesticides in exposure water (i.e. water used for the experiments) attest initiation was reported. In addition, negative controls were not utilized; only solvent (acetone) controls were included. The study authors cite that pesticide application in southern Florida is highest during the dry winter months (December-April), when primary agricultural production occurs, and lightest during the rainy summer months (May-November). However, the sampling periods (October and February) are not necessarily reflective ofthese reported extremes.

There may be little relevance ofthe results from these experiments to natural systems, despite the natural source ofthe communities, given that the microorganisms were incubated in 600 mL beakers containing nutrient amended, filtered (0.22-llm) 

### canal water under unspecified conditions oflight and temperature. During the 7-day colonization period, canal water was not monitored for pH, temperature, turbidity, dissolved oxygen, alkalinity, total nitrogen, total phosphorus, conductivity, andparticulate and dissolved organic matter.
Phytoplankton and zooplankton were not identified at the species level to assess changes in species composition.

This study also examines the potential effects ofchlorothalonil and endosulfan on microbial assemblages collected from the C-111 canal in southernFlorida. Endosulfan was detected inthe canal water at sites C and E at 7.84 ± 3.64 ng/L (5.34-11.97 

### ng/L) and 0.76 ± 0.29 ng/L (0.30- 1.07 ng/L), respectively, in October 1999, and at436 ± 509 ng/L (183-1,350 

### ng/L) and 16.4 ± 8.58 ng/L (7.35-27.5 ng/L), respectively, inFebruary 2000. Canal waterwas also analyzed for chlorothalonil; however, the analytical method did notprovide adequate results. Tables 2 and 3 

### ofthe study were not included inthis review because the statistical analyses pooled responses from atrazine, chlorothalonil, and endosulfanexposure.
Description ofUse in Document: 

### Qualitative

Rationale for Use: 

### The study contributes to the weight ofevidence regarding effects ofatrazine on aquatic plant communities.

Limitations ofStudy: 

### There were multiple deficiencies inthe design, implementation, and reporting ofthis study. No chemical analysis (atrazine or otherpesticides) ofexposure water or the controls was conductedpriorto study initiation. Atrazine and endosulfan were assumed to be present, as they were detected in the canal water during the interval when microbial assemblages were colonizingthe artificial substrates. However, previously documented fluctuations in measuredpesticide concentrations inthe waterbody reinforce the factthatthe sample water (used in experiments) concentrations may nothave been reflective ofthe inconcurrently

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### Tertiary Reviewer: AnitaPease, M.S., Senior Biologist, U.S. EPA measured surface water concentrations. Canal water was notmonitored for pH, temperature, turbidity, alkalinity, total nitrogen, total phosphorus, conductivity, or organic carbon during the colonizationperiod. No negative (solvent-free) controls were included. Incubation conditions (light source, photoperiod, temperature) during the acute exposure experiments were not described. Phytoplankton and zooplankton were not identified atthe species level to assess changes in species composition. Some results were presented as pooled data which combined data from atrazine treatments, exposure durations, sites, and seasons. Finally, for some endpoints, results were not reported for all experiments and exposure durations.


### Primary Reviewer: Lynne Binari, M.S., Subcontractorto Cambridge Environmental Inc.


### Secondary Reviewer: Catherine Aubee, Biologist, Environmental Fate and Effects Division U.S. Environmental Protection Agency**Ct~** 

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