Arch Environ Contam Toxicol (2014) 66:450–462 DOI 10.1007/s00244-014-0002-1

Interactive Effects of Mosquito Control Insecticide Toxicity, Hypoxia, and Increased Carbon Dioxide on Larval and Juvenile Eastern Oysters and Hard Clams R. N. Garcia • K. W. Chung • P. B. Key • L. E. Burnett • L. D. Coen • M. E. DeLorenzo

Received: 10 December 2013 / Accepted: 27 January 2014 / Published online: 16 February 2014 Ó Springer Science+Business Media New York (outside the USA) 2014

Abstract Mosquito control insecticide use in the coastal zone coincides with the habitat and mariculture operations of commercially and ecologically important shellfish species. Few data are available regarding insecticide toxicity to shellfish early life stages, and potential interactions with abiotic stressors, such as low oxygen and increased CO2 (low pH), are less understood. Toxicity was assessed at 4 and 21 days for larval and juvenile stages of the Eastern oyster, Crassostrea virginica, and the hard clam, Mercenaria mercenaria, using two pyrethroids (resmethrin and permethrin), an organophosphate (naled), and a juvenile growth hormone mimic (methoprene). Acute toxicity (4day LC50) values ranged from 1.59 to [10 mg/L. Overall, clams were more susceptible to mosquito control insecticides than oysters. Naled was the most toxic compound in oyster larvae, whereas resmethrin was the most toxic compound in clam larvae. Mortality for both species generally increased with chronic insecticide exposure (21-day LC50 values ranged from 0.60 to 9.49 mg/L). Insecticide exposure also caused sublethal effects, including decreased

Electronic supplementary material The online version of this article (doi:10.1007/s00244-014-0002-1) contains supplementary material, which is available to authorized users. R. N. Garcia
L. E. Burnett Grice Marine Laboratory, College of Charleston, Charleston, SC 29412, USA K. W. Chung
P. B. Key
M. E. DeLorenzo (&) National Ocean Service, National Oceanic and Atmospheric Administration, Charleston, SC 29412, USA e-mail: [email protected] L. D. Coen Department of Biological Sciences and HBOI, Florida Atlantic University, Fort Pierce, FL 34946, USA

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swimming activity after 4 days in larval oysters (4-day EC50 values of 0.60 to 2.33 mg/L) and decreased growth (shell area and weight) in juvenile clams and oysters after 21 days (detected at concentrations ranging from 0.625 to 10 mg/L). Hypoxia, hypercapnia, and a combination of hypoxia and hypercapnia caused mortality in larval clams and increased resmethrin toxicity. These data will benefit both shellfish mariculture operations and environmental resource agencies as they manage the use of mosquito control insecticides near coastal ecosystems.

Insecticides are used to control mosquito populations and prevent the transmission of diseases, such as malaria, dengue fever, and several forms of encephalitis, including West Nile virus (e.g. Hales et al. 2002; Huba´lek and Halouzka 1999; Martens and Hall 2000). Mosquito control insecticides can be classified by the life stage that they target (larvicides or adulticides) and by their chemical class. Adulticides commonly used today consist of two chemical classes, organophosphates (OPs) and pyrethroids. Larvicides include microbial compounds, insect growth-regulating compounds, and a variety of oils and monomolecular films. The present study tested three adulticide chemicals (resmethrin, permethrin, and naled) and one larvicide (methoprene) for their toxicity to clams and oysters. These compounds were selected based on their frequency of use in 14 southeastern United States coastal counties and a lack of toxicity data for estuarine molluscs, particularly early life stages of Mercenaria mercenaria and Crassostrea virginica (DeLorenzo et al. 2012). The test chemicals have various mechanisms of toxicity in target insect species. Resmethrin and permethrin are pyrethroids, which act to disrupt sodium-channel function in neural membranes causing constant stimulation of the neuron

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and ultimately leading to paralysis and death (Miller and Salgado 1985; Soderlund and Bloomquist 1989). Naled, an OP, blocks synaptic nerve transmission by inhibiting acetylcholinesterase (AChE) activity leading to a fatal accumulation of acetylcholine in the synaptic cleft (Habig and DiGiulio 1991). Methoprene is a juvenile growth hormone mimic and acts to disrupt normal insect metamorphosis and molting activity; thus, mosquito larvae are prevented from becoming adult mosquitoes (Henrick 2007). The proximity of residential communities and agricultural areas to aquatic systems increases the potential for overspray, unintentional drift, or runoff of mosquito control insecticides (e.g. Lehotay et al. 1998; Reichenberger et al. 2007), and the presence of these chemicals, even in protected habitats, has been documented (e.g. Hennessey et al. 1992; Pierce et al. 2005). When mosquito control spraying corresponds with low tide, exposed habitats, such as marsh flats and oyster reefs, may receive a more concentrated insecticide application, thus increasing risk to estuarine species, such as juvenile crustaceans and recently settled juvenile bivalves (Bolton-Warberg et al. 2007). Crassostrea virginica and M. mercenaria are two estuarine bivalve species of ecological and economic importance. Both filter suspended particulate matter and sediment, thus increasing light penetration and buffering water flow (Dame 1996; Newell 2004). Crassostrea virginica and M. mercenaria also control phytoplankton dynamics and nutrient processes (Kellogg et al. 2013; Newell 2004) and support a high density and richness of nekton (e.g. Atlantic States Marine Fisheries Commission 2007; Grabowski et al. 2012; Newell 2004; Peterson et al. 2003; Stunz et al. 2010). Bivalve aquaculture also mirrors many of those services previously mentioned deploying both native and nonnative species often in very high densities along the estuarine and marine coasts of the United States (e.g. Coen et al. 2011; Dumbauld et al. 2009). Crassostrea virginica and M. mercenaria are also important prey species for numerous invertebrate and vertebrate species; thus, much effort is currently underway to restore and enhance these bivalve species (Atlantic States Marine Fisheries Commission 2007; Coen et al. 2011; Dumbauld et al. 2009). Commercial shell-fishing represents a large source of income and revenue with the United States domestic oyster and hard clam landings for 2010 estimated at $117,590,000 and $40,886,000, respectively (National Marine Fisheries Service 2011). Shellfish growers, however, are concerned that spraying near their hatcheries may contaminate their facilities and source waters, a concern that is warranted because there is a lack of data on the toxicity of mosquito insecticides for hard clams and Eastern oysters (DeLorenzo et al. 2012). There is also a need for such data to properly manage shellfish populations, which have been decreasing worldwide (Beck et al. 2011). Because younger life stages

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are important for the success of both native and hatchery populations and are generally more sensitive to environmental stress (DeLorenzo and Fulton 2012), larval and juvenile stages should receive more research and management focus. Such information is especially important because insecticide spraying occurs during the summer months (e.g. Milam et al. 2000) coincident with spawning of C. virginica and M. mercenaria in the eastern United States (Kennedy and Krantz 1982; Kraeuter and Castagna 2000). As estuarine species, C. virginica and M. mercenaria experience large and regular fluctuations in temperature, salinity, oxygen, CO2, and pH (Cai and Wang 1998; Kemp 1989; Kraeuter and Castagna 2000; Shumway 1996). For example, pH can range from 6.6 to 8.2, and dissolved oxygen levels can range from 0 to 200 % during a 48-h sampling period in South Carolina tidal creeks (Holland et al. 2004). Similar fluctuations may also occur in mariculture settings, which are usually situated adjacent to estuaries where they use ambient water in hatcheries (e.g. Forrest et al. 2009; Wilson 2002). Physiological stress or chemical changes induced by decreased oxygen and pH may modify the toxicity of insecticides to shellfish. For example, bivalves may increase filtration rate under low oxygen conditions, which may increase insecticide uptake. Similarly, changes in pH can alter chemical degradation rates and bioavailability. Thus, testing the toxicity of mosquito control compounds under hypoxic and hypercapnic (increased CO2) conditions is vital to understanding the risk posed to larval and juvenile molluscs in coastal regions. The objectives of this study were to (1) determine acute toxicity in larval and juvenile clams and oysters of one compound used for larval mosquito control (methoprene) and three compounds used for adult mosquito control (two pyrethroids [permethrin and resmethrin], and one OP [naled]); (2) determine mortality and effects on growth in juvenile clams and oysters after chronic exposure to the same mosquito control compounds; and (3) determine the multistressor effects of the toxicity of one mosquito control compound (resmethrin) on survival in larval clams combined with one of the following: hypoxia, hypercapnia, and a combination of hypoxia and hypercapnia.

Materials and Methods Acute Toxicity Testing Larval Bivalves Newly hatched (24–48 h old) M. mercenaria and C. virginica were purchased from Bay Shellfish Inc. (Terra Ceia, Florida, USA) and Fishers Island Oyster Farm (Fishers Island, New York, USA), respectively. Seawater used in

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the experiments was supplied to the laboratory from the Charleston Harbor estuary: filtered (5 lm), ultravioletsterilized, filtered with activated carbon, and diluted with deionized water. Larvae were acclimated to laboratory conditions (16:8-h light-to-dark cycle, 25 °C, 20 ppt seawater) during a 48- to 72-h period in glass finger bowls (2 L) with gentle aeration. Larvae were fed approximately 20,000 cells/mL of the algae Isochrysis galbana (also supplied by Bay Shellfish) daily. Larval testing was performed in 24-well polystyrene plates coated with hydrogel (Corning, Lowell, MA, USA) to decrease adherence of insecticides (Chandler et al. 2004). One free-swimming larva (7–9 days old) was placed in each well with 2 mL of test solution and 12,000 cells/mL of I. galbana. For each chemical tested (resmethrin, permethrin, naled, and methoprene), the acute tests included a control and five nominal insecticide concentrations (0.12, 0.37, 1.11, 3.33, and 10 mg/L) with three replicate 24-well plates/ concentration (total of 18 well plates/test chemical; n = 72 individuals/treatment). The concentrations were determined using range finder tests. The plates were placed on an orbital shaker (100 rpm) in a Percival environmental chamber set to the above-mentioned laboratory conditions. For the larval tests, the seawater was further filtered to 0.22 lm. Insecticide stock solutions were prepared using technical-grade material ([97 % purity, Sigma-Aldrich, St. Louis, Missouri, USA) dissolved in 100 % high-performance liquid chromatography–grade acetone, and all treatments and controls had a final acetone concentration of 0.1 %. It was previously determined that the acetone carrier solvent at concentrations B0.1 % was not found to produce any significant effects on larval or juvenile bivalves compared with seawater controls (data not shown). Larval mortality was determined daily by visual inspection with a dissecting microscope. At each daily treatment renewal, surviving larvae were transferred to new 24-well plates with new insecticide solution and fed. Water quality parameters (salinity, temperature, oxygen, and pH) were measured daily from the 24-h-old test solutions. After 96 h, mortality (the number of dead larvae out of 24 on each plate) was assessed, and median lethal (LC50) and LC10 concentrations with 95 % confidence intervals (CIs) were determined using Probit Analysis (SAS V.9.1.3, SAS Institute, Inc., Cary, NC, USA). In some cases, the data were log-transformed. For each experiment, the no observed-effect (NOEC) and lowest observed effect concentrations (LOEC) were identified using one-way analysis of variance (ANOVA) with Dunnett’s test (Zar 1999). During testing, we noted a decrease in the number of oyster larvae swimming with insecticide treatment compared with control larvae. Swimming activity was determined by visual observation of individual oysters in the well plate using the dissecting microscope. Any swimming

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activity was recorded as swimming, whereas no swimming activity in a 15-second observation period was recorded as non-swimming. A swimming 96-h EC50 value was calculated for larval oysters using the linear interpolation method for sublethal toxicity (Norberg-King 1993). Juvenile Bivalves Mercenaria mercenaria and C. virginica juveniles (approximately 1 mm after settlement) were supplied by Bay Shellfish Inc. and Fishers Island Oyster Farm, respectively. Juveniles were transferred to glass finger bowls and held under the above-described laboratory conditions. Acute testing for juvenile clams and oysters were completed in a similar manner as described previously for the larvae. The tests included a control and five nominal insecticide concentrations (0.63, 1.25, 2.5, 5, and 10 mg/L), and concentrations were determined using range-finder tests. Tests were completed in wide-mouth (473 mL) glass jars with 180 mL of a given test solution per jar. Thirty juvenile bivalves were added to each jar (n = 3 replicate jars/ treatment). Juvenile bivalves were not fed during the acute test. Mortality (the number of dead out of 30 individuals in each jar) was determined at 96 h, and LC50 and LC10 values were estimated using probit analysis. NOEC and LOEC were assessed using ANOVA followed by Dunnett’s test. Significant differences (p \ 0.05) between LC50 values for different species, life stages, and test chemicals were determined using the LC50 ratio test (Wheeler et al. 2006). Chronic Toxicity Juvenile bivalves were also tested in a chronic (21-day) exposure to determine mortality and any sublethal effects on growth. Chronic testing followed the same protocols and included the same concentrations used for the acute testing with juveniles except that the test media were renewed every 72 h and the animals fed I. galbana at a concentration of 20,000 cells/mL after each renewal. After 21 days, mortality and growth (shell area and dry weight) were assessed. LC50, LC10, NOEC, and LOEC values were determined. Dead animals were excluded from sublethal end-point analysis. The mean dry mass (mg) of pooled individuals from each replicate was determined by drying overnight at 68 °C. Individual shell area (mm2) measurements were made using an Olympus dissecting microscope with digital camera and Image Pro Plus 6.3 software (Media Cybernetics, Inc., Rockville, MD, USA). One-way ANOVA with Dunnett’s test was used to determine whether the shell measurements and dry weights of the treatment organisms were different from those of the control organisms within each test. Where possible, a 21-day EC50 value for growth was determined.

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Multistressor Tests on Survival Acute, 96-h static renewal testing of insecticide exposure and abiotic variables [hypoxia and hypercapnia (increased CO2)] was performed using only larval clams because they could be obtained year-round. Resmethrin was selected for multistressor testing based on results of the acute and chronic exposures. A two-factorial design was set up to test both resmethrin concentration and level of oxygen and carbon dioxide. Three separate experiments were performed: (1) resmethrin with hypoxia, (2) resmethrin with hypercapnia, (3) resmethrin with hypercapnic hypoxia. Control conditions in each experiment were generated with laboratory-supplied air to yield normoxia (approximately 6 mg/L and 70 % air saturation) and nomocapnia (approximately pH 9), which were based on average measurements obtained during acute testing with wellaerated seawater. Three treatments were bubbled with a combination of 4 % oxygen and 96 % nitrogen for hypoxia (approximately 2 mg/L O2/L or 20 % air saturation); a combination of 19 % oxygen (near air saturation), 79 % nitrogen, and 2 % CO2 for hypercapnia (approximately pH 7); and a combination of 4 % oxygen (20 % air saturation), 94 % nitrogen, and 2 % CO2 for hypercapnic hypoxia (Cochran and Burnett 1996). Gas mixtures were generated using a gas mixer (Pegas 4000 MF; Columbus Instruments). Both carbon dioxide treatments (2 % CO2) resulted in a pH of approximately 7, whereas the normoxic and hypoxic treatments without CO2 had a pH of approximately 8. These levels of oxygen and CO2 were chosen because they represent values commonly seen in estuarine and especially tidal creek habitats of the test organisms used in this study (Cochran and Burnett 1996; Lerberg et al. 2000). Each experiment included a control (no resmethrin), 0.79 mg/L resmethrin (half of the LC50 value determined for larval clams in acute testing), and 1.59 mg/L resmethrin (the LC50 value determined); nominal concentrations. Each treatment had three replicate 125-mL Erlenmeyer flasks fitted with silicon stoppers with ports for gas control. Each flask contained 24 larvae and 100 mL of test solution. Test solutions were renewed and water quality parameters measured every 24 h. Mortality was determined at 96 h and a two-way ANOVA (SAS V.9.1.3) was used to determine significant differences (p \ 0.05) in mortality due to the individual factors and interaction between the two factors (Zar 1999). Environmental Hazard Assessment The data generated from the acute toxicity experiments were used to prepare an environmental hazard assessment specifically for clams and oysters and the mosquito control insecticides tested. Application data [kg of active

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ingredient (AI)/ha] were obtained from the pesticide product labels, which were then used to calculate the maximum estimated environmental (EEC) concentration for surface water. This assumes that the product is applied directly to a 1-ha pond (30 cm depth) at the application rate and is well mixed. It does not take into account wind, tidal stage, etc. A hazard quotient was calculated for each chemical as the estimated water concentration/96-h LC50 value for the most sensitive species and life stage (Suter 1995).

Results Acute Effects of Mosquito Control Insecticides on Clam and Oyster Survival All water quality parameters were within acceptable ranges for larval clam and oyster acute tests (Table 1). In larval clams, only the highest concentration tested (10 mg/L) for naled (p = 0.014) and permethrin (p = 0.0019) caused significant mortality compared with the controls (one-way ANOVA and Dunnett’s test) [Supplementary (Suppl.) Fig. 1]. At 10 mg/L, naled caused 53 % mortality, and permethrin caused 60 % mortality. Resmethrin concentrations C1.11 mg/L caused significantly greater mortality than the control treatment (p \ 0.0001) with 40 % mortality at 1.11 mg/L, 63 % at 3.33 mg/L, and 74 % at 10 mg/L. Methoprene concentrations C1.11 mg/L caused significantly greater mortality than the control treatment (p \ 0.0001) with 17 % mortality at 1.11 mg/L, 72 % mortality at 3.33 mg/L, and 81 % mortality at 10 mg/L. The 96-h LC50 values determined in larval clams were 1.59 mg/L for resmethrin, 7.65 mg/L for permethrin, 8.73 mg/L for naled, and 2.81 mg/L for methoprene (Table 2). The 96-h LC10 values determined in larval clams for resmethrin, permethrin, and naled were less than the that of the lowest concentration tested (0.12 mg/L) and for methoprene was 0.7 mg/L (Table 2). For larval oysters, naled, permethrin, and resmethrin concentrations C3.33 mg/L caused significantly greater mortality than the control (all p values \0.0001; one-way ANOVA and Dunnett’s test) (Suppl. Fig. 2). Naled exposure caused 38 % mortality at 3.33 mg/L and 57 % mortality at 10 mg/L. Permethrin and resmethrin exposure both caused 14 % mortality at 3.33 mg/L and approximately 32 % mortality at 10 mg/L. Mortality at the highest methoprene concentration tested (10 mg/L) was 11 %. The 96-h LC50 values determined in larval oysters were [10 mg/L (the highest concentration tested) for resmethrin, permethrin, and methoprene and 8.26 mg/L for naled (Table 3). The 96-h LC10 values determined in larval oysters were 1.89 mg/L for naled, 4.86 mg/L for

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Table 1 Summary of water quality parameters for each acute and chronic toxicity test Species

Life stage

Insecticide

Exposure

Temp (°C)

Salinity (ppt)

DO (mg/L)

pH

M. mercenaria

Larval

Resmethrin

Acute

21.8 ± 0.03

20.5 ± 0.02

6.66 ± 0.010

8.02 ± 0.003

M. mercenaria

Larval

Permethrin

Acute

21.6 ± 0.11

20.6 ± 0.02

6.73 ± 0.028

7.98 ± 0.009

M. mercenaria

Larval

Naled

Acute

22.0 ± 0.03

20.2 ± 0.01

6.67 ± 0.01

8.01 ± 0.004

M. mercenaria

Larval

Methoprene

Acute

22.6 ± 0.08

21.7 ± 0.02

7.27 ± 0.015

7.85 ± 0.009

M. mercenaria

Juvenile

Resmethrin

Acute

23.6 ± 0.02

20.4 ± 0.02

6.96 ± 0.006

7.87 ± 0.002

M. mercenaria

Juvenile

Permethrin

Acute

23.7 ± 0.02

20.4 ± 0.02

6.93 ± 0.006

7.92 ± 0.001

M. mercenaria

Juvenile

Naled

Acute

22.7 ± 0.04

20.6 ± 0.02

6.88 ± 0.010

7.94 ± 0.003

M. mercenaria

Juvenile

Methoprene

Acute

23.0 ± 0.14

21.0 ± 0.01

7.21 ± 0.048

7.99 ± 0.008

M. mercenaria

Juvenile

Resmethrin

Chronic

23.3 ± 0.01

20.2 ± 0.01

6.58 ± 0.008

7.94 ± 0.002

M. mercenaria

Juvenile

Permethrin

Chronic

23.7 ± 0.01

20.6 ± 0.01

6.76 ± 0.004

7.93 ± 0.002

M. mercenaria M. mercenaria

Juvenile Juvenile

Naled Methoprene

Chronic Chronic

23.3 ± 0.01 22.9 ± 0.06

20.8 ± 0.01 21.0 ± 0.01

6.69 ± 0.005 6.92 ± 0.046

7.93 ± 0.001 8.02 ± 0.008

C. virginica

Larval

Resmethrin

Acute

21.4 ± 0.02

21.7 ± 0.02

7.23 ± 0.004

7.93 ± 0.001

C. virginica

Larval

Permethrin

Acute

21.5 ± 0.03

21.0 ± 0.01

7.22 ± 0.005

7.92 ± 0.001

C. virginica

Larval

Naled

Acute

21.7 ± 0.02

21.5 ± 0.03

7.26 ± 0.005

7.85 ± 0.002

C. virginica

Larval

Methoprene

Acute

22.6 ± 0.08

21.7 ± 0.02

7.27 ± 0.015

7.85 ± 0.009

C. virginica

Juvenile

Resmethrin

Acute

23.0 ± 0.04

20.7 ± 0.02

6.99 ± 0.010

8.10 ± 0.002

C. virginica

Juvenile

Permethrin

Acute

22.9 ± 0.03

20.5 ± 0.02

7.00 ± 0.010

8.03 ± 0.001

C. virginica

Juvenile

Naled

Acute

23.3 ± 0.03

20.5 ± 0.02

7.04 ± 0.009

8.04 ± 0.002

C. virginica

Juvenile

Methoprene

Acute

23.9 ± 0.09

20.5 ± 0.01

7.02 ± 0.033

8.04 ± 0.002

C. virginica

Juvenile

Resmethrin

Chronic

23.3 ± 0.01

20.5 ± 0.01

6.93 ± 0.004

8.11 ± 0.001

C. virginica

Juvenile

Permethrin

Chronic

23.0 ± 0.01

20.7 ± 0.01

6.99 ± 0.004

8.06 ± 0.001

C. virginica

Juvenile

Naled

Chronic

23.4 ± 0.01

20.8 ± 0.01

6.97 ± 0.003

8.02 ± 0.002

C. virginica

Juvenile

Methoprene

Chronic

23.1 ± 0.07

21.0 ± 0.01

7.16 ± 0.042

8.10 ± 0.020

All tests were completed under 16:8-h light-to-dark cycle. Values are average ±SEM of the daily measurements taken in each test Table 2 Summary of probit analysis results for acute (4-day exposure) and chronic (21-day exposure) effects of resmethrin, permethrin, naled, and methoprene on survival of larval and juvenile clams, M. mercenaria Life stage

Insecticide

Exposure

LC50 (95 % CI) (mg/L)

LC10 (95 % CI) (mg/L)

NOEC (mg/L)

LOEC (mg/L)

Larval

Resmethrin

Acute

1.59 (0.94–2.40)

\0.12

0.37

1.11

Larval

Permethrin

Acute

7.65 (6.18–10.03)

\0.12

3.33

10

\0.12

Larval

Naled

Acute

8.73 (6.70–12.70)

3.33

10

Larval

Methoprene

Acute

2.81 (2.06–3.93)

0.70 (0.36–1.05)

0.37

1.11

Juvenile

Resmethrin

Acute

8.24 (7.36–9.39)

1.67 (0.69–2.45)

1.25

2.5

Juvenile

Permethrin

Acute

9.10 (7.53–11.77)

\0.63

0.63

1.25

Juvenile Juvenile

Naled Methoprene

Acute Acute

4.54 (2.29–27.34) [10

\0.63 [10

\0.63 10

0.63 [10

Juvenile

Resmethrin

Chronic

7.78 (5.66–13.07)

\0.63

2.5

Juvenile

Permethrin

Chronic

0.60 (0.01–1.41)

\0.63

\0.63

0.63

Juvenile

Naled

Chronic

2.82 (1.34–7.00)

\0.63

\0.63

0.63

Juvenile

Methoprene

Chronic

0.68 (0.28–1.03)

\0.63

\0.63

0.63

resmethrin, 4.95 mg/L for permethrin, and 8.17 mg/L for methoprene (Table 3). All water quality parameters were within acceptable ranges for acute tests with juvenile clams and oysters (Table 1). In juvenile clams, acute exposures to resmethrin, permethrin, and naled caused significant mortality

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compared with the controls (p values \ 0.0001, one-way ANOVA and Dunnett’s test) (Suppl. Fig. 3). Naled caused significant mortality at all concentrations tested ranging from 38 % mortality at 0.63 mg/L to 52 % mortality at 10 mg/L. Permethrin treatments C1.25 mg/L had significantly greater mortality than the control ranging from 17 %

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Table 3 Summary of probit analysis results for acute (4-day exposure) and chronic (21-day exposure) effects of resmethrin, permethrin, naled, and methoprene on survival of larval and juvenile oysters, C. virginica Life stage

Insecticide

Exposure

LC50 (95 % CI) (mg/L)

LC10 (95 % CI) (mg/L)

NOEC (mg/L)

LOEC (mg/L)

Larval

Resmethrin

Acute

[10

4.86 (3.67–5.97)

1.11

3.33

Larval

Permethrin

Acute

[10

4.95 (3.73–6.08)

1.11

3.33

Larval

Naled

Acute

8.26 (6.52–11.31)

1.89 (–0.31 to 3.41)

1.11

3.33

Larval

Methoprene

Acute

[10

8.17 (5.05–20.93)

10

[10

Juvenile

Resmethrin

Acute

[10

3.94 (–)

10

[10

Juvenile

Permethrin

Acute

[10

2.79 (–)

10

[10

Juvenile Juvenile

Naled Methoprene

Acute Acute

7.84 (7.01–8.69) [10

3.35 (1.71–4.46) 2.79 (–)

2.5 10

5 [10

Juvenile

Resmethrin

Chronic

9.49 (8.27–11.26)

\0.63

5

10

Juvenile

Permethrin

Chronic

4.21 (3.46–5.16)

\0.63

1.25

2.5

Juvenile

Naled

Chronic

1.14 (0.43–1.81)

\0.63

\0.63

0.63

Juvenile

Methoprene

Chronic

1.32 (0.98–1.68)

\0.63

\0.63

0.63

mortality at 1.25 mg/L to 51 % at 10 mg/L. Resmethrin treatments C2.5 mg/L caused significantly greater mortality than the control with 14 % mortality at 2.5 mg/L, 32 % mortality at 5 mg/L and 60 % mortality at 10 mg/L. Acute exposure to methoprene did not cause significant mortality in juvenile clams at any of the concentrations tested. The 96-h LC50 values determined in juvenile clams were 8.24 mg/L for resmethrin, 9.10 mg/L for permethrin, 4.54 mg/L for naled, and [10 mg/L for methoprene (Table 2). The 96-h LC10 values determined in juvenile clams were [10 mg/L for methoprene, 1.67 mg/L for resmethrin, and \0.63 mg/L for permethrin and naled (Table 2). For juvenile oysters, there was a significant increase in mortality versus the control for naled concentrations C5 mg/L (p \ 0.0001, one-way ANOVA and Dunnett’s test) (Suppl. Fig. 4). Naled exposure caused 24 % mortality at 5 mg/L and 81 % mortality at 10 mg/L. No significant mortality occurred with acute exposure to permethrin (p = 0.16), resmethrin (p = 0.20), or methoprene (p = 0.7342) at any of the concentrations tested. At the highest concentration tested (10 mg/L), mortality was 18 % in the permethrin exposure and 7 % in the resmethrin and methoprene exposures. The 96-h LC50 values determined in juvenile oysters were [10 mg/L (the highest concentration tested) for resmethrin, permethrin, and methoprene and 7.84 mg/L for naled (Table 3). The 96-h LC10 values determined in juvenile oysters were 3.94 mg/L for resmethrin, 2.79 mg/L for permethrin, 2.79 mg/L for methoprene, and 3.35 mg/L for naled (Table 3).

increased length of exposure to mosquito control insecticides led to an increase in toxicity to juvenile clams (Table 2) and oysters (Table 3). Chronic exposure to permethrin increased toxicity (decreasing the LC50 value) approximately 15-fold from 9.10 mg/L to 0.60 mg/L in clams and from [10 mg/L to 4.21 mg/L in oysters. Significant chronic effects of permethrin on mortality were seen at concentrations C0.625 mg/L in clams (Suppl. Fig. 5) and C2.5 mg/L in oysters (Suppl. Fig. 6). Methoprene also exhibited a significant increase in toxicity with chronic exposure with clam LC50 decreasing from [10 mg/L to 0.68 mg/L and oyster LC50 decreasing from [10 mg/L to 1.32 mg/L. Significant effects of methoprene on mortality were seen at concentrations C0.625 mg/L in clams (Suppl. Fig. 5) and oysters (Suppl. Fig. 6). Chronic exposure to naled slightly increased toxicity in clams (decreasing LC50 from 4.54 mg/L to 2.82 mg/L) and caused an approximately 7-fold increase in toxicity in oysters (decreasing LC50 from 7.84 mg/L to 1.14 mg/L). Significant effects of naled on mortality were seen at concentrations C0.625 mg/L in clams (Suppl. Fig. 5) and oysters (Suppl. Fig. 6). Acute resmethrin LC50 values for clams (8.24 mg/L) and oysters ([10 mg/L) were similar to chronic LC50 values (clams = 7.78 mg/L; oysters = 9.49 mg/L). Significant effects of resmethrin on mortality were seen at concentrations C5 mg/L in clams (Suppl. Fig. 5) and C10 mg/L in oysters (Suppl. Fig. 6). Sublethal Effects of Mosquito Control Insecticides on Clams and Oysters Swimming Activity in Larval Oysters After Acute Exposure

Chronic Effects of Mosquito Control Insecticides on Clam and Oyster Survival All water quality parameters were within acceptable ranges for larval clam and oyster chronic tests (Table 1). In general,

The number of larval oysters swimming after 96 h was significantly decreased by exposure to permethrin (p = 0.0004) and methoprene (p = 0.0013) at concentrations C3.33 mg/L (Fig. 1). Naled (p \ 0.0001) and

123

456

Arch Environ Contam Toxicol (2014) 66:450–462 naled

0.05

resmethrin 0.04

15

10

* *

5

*

* *

*

0 Control

0.12

0.37

1.11

* **

3.33

permethrin

0.045

Clam dry weight (mg)

Average number of oysters swimming

methoprene naled methoprene permethrin resmethrin

20

*

10

0.035 0.03 0.025 0.02 0.015 0.01 0.005

Insecticide concentration (mg/L) 0

Fig. 1 Effect of insecticide exposure on number of larval oysters swimming after 4-day acute exposures. Data are means of three replicate treatments (24 oysters each) ±SEM for each treatment. Asterisks indicate treatments that were significantly different from the respective control for each insecticide (one-way ANOVA, Dunnett’s test, p \ 0.05)

naled methoprene permethrin resmethrin

3.5

Control

0.625

1.25

2.5

5

10

Insecticide concentration (mg/L)

Fig. 3 Effect of insecticide exposure on juvenile clam dry weight after 21 days. Data are means of three replicate treatments ±SEM for each treatment. Clams within each replicate were pooled for dry weight. Asterisks indicate treatments that were significantly different from the respective control for each insecticide (one-way ANOVA, Dunnett’s test, p \ 0.05)

0.60–2.22 mg/L) for methoprene, and 2.33 mg/L (95 % CI 1.52–3.52 mg/L) for permethrin.

Clam shell area (mm2)

3

Growth in Juvenile Clams and Oysters After Chronic Exposure

2.5

*

2

* *

*

*

1.5

*

* *

1

0.5

0 Control

0.625

1.25

2.5

5

10

Insecticide concentration (mg/L)

Fig. 2 Effect of insecticide exposure on juvenile clam shell area after 21-day exposure. Data are means of three replicate treatments (30 clams each) ±SEM for each treatment. Asterisks indicate treatments that were significantly different from the respective control for each insecticide (one-way ANOVA, Dunnett’s test, p \ 0.05)

resmethrin (p = 0.0001) significantly decreased the number of oysters swimming at concentrations C1.11 mg/L (Fig. 1). The 96-h EC50 values determined for the effect on swimming in larval oysters were: 0.60 mg/L (95 % CI 0.31–0.74 mg/L) for naled, 0.93 mg/L (95 % CI 0.37–1.55 mg/L) for resmethrin, 0.99 mg/L (95 % CI

123

Juvenile clam and oyster growth, as measured by shell area, was also affected by chronic exposure to most of the insecticides tested. Resmethrin did not affect clam shell area (p = 0.0665) (Fig. 2). Clam growth was significantly inhibited by permethrin concentrations C5 mg/L (p = 0.0002) and naled concentrations C2.5 mg/L (p \ 0.0001) (Fig. 2). The effect of methoprene on clam growth was variable with a significant effect detected (p \ 0.0001), but this occurred only at concentrations of 0.625, 1.25, and 10 mg/L (Fig. 2). A [50 % effect on clam shell area was not observed with any of the insecticides tested. Mean clam weight was significantly less than controls at all permethrin (p \ 0.0001) and methoprene (p = 0.0055) concentrations tested and at naled concentrations C1.25 mg/ L (p = 0.0032) (Fig. 3). Resmethrin significantly decreased mean clam weight (p = 0.0230) but did so only at the highest concentration (10 mg/L) (Fig. 3). A 21-day EC50 value for mean clam weight of 1.21 mg/L (95 % CI 0.60–2.23) was calculated for naled, 0.91 mg/L for methoprene (95 % CI 0.63–1.06), and 0.63 mg/L (95 % CI 0.53–0.87) for permethrin. Oyster shell area was significantly smaller than the control at all resmethrin (p \ 0.0001) and permethrin

Arch Environ Contam Toxicol (2014) 66:450–462 3

457

Effects of Hypoxia and Resmethrin

naled methoprene permethrin resmethrin

2.5

100 90

‡

Normoxia

*

€

Hypoxia

Percent Mortality

Oyster shell area (mm2)

80

2

1.5

70

‡

60

*

50

‡

40

‡

*

30

1 20 10

0.5

0 CTL

0 Control

0.625

1.25

2.5

5

10

Insecticide concentration (mg/L)

Fig. 4 Effect of insecticide exposure on juvenile oyster shell area after 21-day exposure. Data are means of three replicate treatments (30 oysters each) ±SEM for each treatment. Asterisks indicate treatments that were significantly different from the respective control for each insecticide (one-way ANOVA, Dunnett’s test, p \ 0.05). There were no shell area measurements for naled at 10 mg/L due to 100 % oyster mortality at that concentration

0.025

naled methoprene permethrin resmethrin

0.02

Oyster dry weight (mg,)

0.79

1.59

Resmethrin Concentration (mg/L)

0.015

Fig. 6 Mortality due to hypoxia and resmethrin in larval clams during the multistressor experiment (4-day exposure). Bars are means of three replicate treatments (24 clams each) ±SEM. There was a significant increase in mortality due to resmethrin concentration (àp \ 0.0001) and a significant increase in mortality due to oxygen level (*p \ 0.0001). There was also a significant interaction between resmethrin concentration and oxygen level (€p \ 0.0001) (two-way ANOVA)

Resmethrin concentrations C1.25 mg/L significantly decreased mean oyster weight (p \ 0.0001). Permethrin (p \ 0.0001), methoprene (p \ 0.0001), and naled (p = 0.0004), significantly decreased mean oyster weight at all concentrations tested compared with the control (Fig. 5). The 21-day EC50 value for mean oyster weight was 0.59 mg/L (95 % CI 0.39–0.96) for methoprene, 0.63 mg/L (95 % CI 0.51–1.01) for naled, and 4.13 mg/L (95 % CI 3.33–5.21) for permethrin.

0.01

Comparisons of Toxicity Between Species and Life Stages

0.005

0 Control

0.625

1.25

2.5

5

10

Insecticide concentration (mg/L)

Fig. 5 Effect of insecticide exposure on juvenile oyster dry weight after 21 days. Data are means of three replicate treatments ± SEM for each treatment. Oysters within each replicate were pooled for dry weight. Asterisks indicate treatments that were significantly different from the respective control for each insecticide (one-way ANOVA, Dunnett’s test, p \ 0.05). There were no dry weight measurements for naled at 10 mg/L due to 100 % oyster mortality at that concentration

(p \ 0.0001) concentrations tested and at all methoprene (p \ 0.001) concentrations except for 5 mg/L (Fig. 4). Oysters under naled concentrations C2.5 mg/L had significantly smaller shell area than the control (p \ 0.0001) (Fig. 4). A [50 % effect on oyster shell area was not observed with any of the insecticides tested.

Differences in acute toxicity between species and life stages were determined using LC50 ratio tests. Larval clams were approximately fivefold more sensitive to resmethrin (p = 0.0002) and methoprene (p = 0.0017) than juvenile clams based on 96-h LC50 values. There were no significant differences in LC50 value between larval and juvenile clams for permethrin (p = 0.256) or naled (p = 0.419). Larval clams were sixfold more sensitive to resmethrin (p \ 0.0001) and 4-fold more sensitive to methoprene (p = 0.0004) than larval oysters. There was no significant difference between larval clams and larval oysters for naled (p = 0.909) or permethrin (p = 0.074). There were no significant differences in LC50 between larval and juvenile oysters for any of the insecticides tested (p values [0.05). There were also no significant differences in the LC50 values between juvenile clams and juvenile oysters for any of the insecticides tested (p values [0.05).

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Arch Environ Contam Toxicol (2014) 66:450–462

Effects of Hypercapnia and Resmethrin

100

Normocapnia

90

Hypercapnia

Percent Mortality

80

‡

*

70 60

‡

*

50

‡

*

40

‡

30

compared with the control under normoxia. In the hypercapnic test, there was a significant difference in mortality due to resmethrin concentration (p \ 0.0001) and hypercapnia (p \ 0.0001, Fig. 7). However, there was not a significant interaction between resmethrin concentration and hypercapnia (p = 0.92). In the hypercapnic hypoxic test, there was a significant difference in mortality due to resmethrin concentration (p \ 0.0001) and oxygen and hypercapnic hypoxia (p \ 0.0001, Fig. 8). There was no significant interaction between resmethrin concentration and oxygen and hypercapnic hypoxia (p = 0.084).

20

Hazard Assessment

10 0 CTL

0.79

1.59

Resmethrin Concentration (mg/L)

Fig. 7 Mortality due to hypercapnia and resmethrin in larval clams during the multistressor experiment (4-day exposure). Bars are means of three replicate treatments (24 clams each) ±SEM. There was a significant increase in mortality due to resmethrin concentration (àp \ 0.0001) and a significant increase in mortality due to pH level (*p \ 0.0001). There was no significant interaction detected between pH level and resmethrin concentration (p = 0.92) (two-way ANOVA)

Effects of Hypercapnic Hypoxia and Resmethrin 100

Normocapnic Normoxia

The estimated maximum surface water concentrations were 0.00267 mg/L for resmethrin and permethrin and approximately 0.04 mg/L for naled and methoprene (Table 4). The lowest acute LC50 values used in the assessment were for larval clams except for naled where the juvenile clam value was used. The lowest estimated hazard quotient calculated was for permethrin (0.000349) and the highest hazard quotient calculated was for methoprene (0.0174) (Table 4).

Discussion

90

Percent Mortality

80

Hypercapnic Hypoxia

‡

*

70 60

‡

50 40

*

30

‡ ‡

*

20 10 0 C TL

0.79

1.59

Resmethrin Concentration (mg/L)

Fig. 8 Mortality due to hypercapnic hypoxia and resmethrin treatments in larval clams during the multistressor experiment (4-day exposure). Bars are means of three replicate treatments (24 clams each) ±SEM. There was a significant increase in mortality due to resmethrin concentration (àp \ 0.0001) and a significant increase in mortality due to oxygen and pH level (*p \ 0.0001). There was no significant interaction detected between oxygen and pH

Multistressor Effects on Survival Larval clams had significant mortality due to oxygen level (p \ 0.0001) and resmethrin concentration (p \ 0.0001, Fig. 6). There was also an interaction between oxygen level and resmethrin concentration (p \ 0.0001) with hypoxia causing an 8-fold increase in mortality at 1.59 mg/L

123

This study characterized the toxicity of four mosquito insecticide compounds to the larval and juvenile stages of two important bivalve species, C. virginica and M. mercenaria, and the effects of low oxygen (hypoxia) and low pH due to increased carbon dioxide (hypercapnia) on resmethrin toxicity in larval M. mercenaria. Both of these commercially important species lack data on insecticide toxicity, especially at younger life stages, and both species are vulnerable to effects from hypoxia and hypercapnia (e.g. Dickinson et al. 2012, 2013; Tomanek et al. 2011; Waldbusser et al. 2011). Differences in toxicity were observed among chemicals, species, and life history stages. In general, hard clams were more acutely sensitive than oysters to the insecticides tested. Resmethrin was the most toxic chemical to larval clams, whereas naled was the most toxic chemical tested to juvenile clams and both oyster life history stages. Because juvenile clams are mobile and have siphons that detect light and other environmental cues, they are often envisioned as having more complex sensory systems, which may increase chemical sensitivity (Carriker 2001; Bondarenko et al. 2006; Jaffe 1991). There may also be differences between the two species in how their nervous systems develop, and therefore, how they respond to OPs (AChE target) and pyrethroids (sodium-channel target). Adult M. mercenaria hearts were 10,000 times more sensitive to AChE depression in vitro than C. virginica

Arch Environ Contam Toxicol (2014) 66:450–462

459

Table 4 Hazard assessment of mosquito control insecticides for M. mercenaria and C. virginica Insecticide

Application rate (kg AI/ha)

Mosquito control product

Surface water EEC (mg/L)a

Lowest 96-h LC50 (mg/L)

Species and life stage

Hazard quotient

Resmethrin

0.008

ScourgeÒ 18?54

2.67 9 10-3

1.59

Larval clams

1.68 9 10-3

-3

7.65

Larval clams

3.49 9 10-4

-2

4.54

Juvenile clams

8.26 9 10-3

-2

2.81

Larval clams

1.74 9 10-2

Permethrin Naled Methoprene a

Ò

0.008

Permanone 30-30

0.113

Ò

0.146

Trumpet EC Ò

Altosid

2.67 9 10

3.75 9 10

4.88 9 10

Estimated maximum potential concentration in 30 cm (12 in) of surface water, assumes a 1 ha pond (100 9 100 m2), 30 cm depth

hearts despite the fact that AChE activity recovery in oysters was 100 times greater than that in clams (Roop and Greenberg 1967). Additional research is needed, particularly on early life stages of these species, to elucidate the specific mechanisms of insecticide toxicity to clams and oysters. Differences in sensitivity were also noted between larval and juvenile life stages. In general, larvae were more sensitive to insecticide exposure than juveniles. This trend is consistent with other estuarine species, such as grass shrimp, whose earlier life stages had greater sensitivity to permethrin (DeLorenzo and Fulton 2012) and resmethrin (Key et al. 2005). Differences in chemical toxicity between early and later life history stages may be due to developmental differences in nervous system target sites; for example, AChE activity has been shown to vary significantly by life stage in the grass shrimp (Hoguet and Key 2007). In larval oysters, a sublethal response was noted with an observed decrease in number of larval oysters swimming for all insecticides tested at concentrations approximately tenfold lower than LC50 values for either larvae or juveniles. These results suggest greater sensitivity to insecticides in the earlier life stages and thus a greater potential for negative impacts early on in the species’ recruitment and survival. Bivalve larvae that cannot swim may be forced to settle in unsuitable habitats or they may be unable to settle at all, leading to increased mortality and ultimately a reduction in population recruitment and success (North et al. 2008; Tamburri et al. 1992). Prolonged insecticide exposure increased the toxicity of most of the mosquito control compounds tested for both juvenile clams and oysters. Lower concentrations of all compounds were needed to cause bivalve mortality in the chronic exposures. Bivalve molluscs are capable of closing their shells for days at a time to endure poor water quality conditions (Coen and Heck 1991), which may in part explain the increased toxicity after the 21-day exposure compared with the 4-day exposure. Chronic insecticide exposure also resulted in decreased clam and oyster growth at concentrations lower than the LC50. Decreased growth could be detrimental to both shellfish mariculture production and the health of natural shellfish populations (Coen

and Lukenbach 2000; Luckenbach et al. 2005; Naylor et al. 2000). Physiological stress or chemical changes induced by environmental factors may modify the toxicity of insecticides to aquatic species. The results of the present study showed that decreases in oxygen and increases in CO2, combined with resmethrin exposure, caused a significant increase in mortality in larval clams compared with standard testing conditions. Although estuarine organisms are adapted to survive dramatic fluctuations in oxygen and pH that regularly occur in their habitat (Dickinson et al. 2012, 2013; Tomanek et al. 2011), they may be physiologically challenged by the combined stress of chemical contamination and exposure to hypoxic or hypercapnic conditions. For example, bivalves may increase filtration rates under low oxygen conditions, thus leaving less energy to launch cellular enzyme defenses against chemical contaminants. In addition to the quantified effects on survival, sublethal changes were qualitatively noted under hypoxic and hypercapnic stress. Larval clams exposed to hypoxia were observed to be less mobile after 24 h (stationary and on the bottom of the chambers), whereas the larvae in the normoxic treatment were freely swimming. The decreased activity may impact larval settlement, and as seen in a study by Baker and Mann (1992) at similar oxygen levels, settlement rates were reduced in larval clams exposed to hypoxia. Clam larvae exposed to hypercapnia appeared to have thinner shells with irregular edges. Decreases in shell formation or decalcification at decreased pH have been seen in other calcifying organisms (e.g. Dove and Sammut 2007; Kuffner et al. 2008; Kurihara et al. 2007; Miller et al. 2009; Watson et al. 2009). Prolonged hypoxic and hypercapnic conditions will be detrimental to hard clam early life stages leading to physiological stress and abnormal development. Future degradation of water quality conditions due to pollution or climate change may increase oxygen and pH stress to both wild bivalve populations and mariculture operations. Overall, the results of this study suggest that the likelihood of adverse effects to early life stages of clams and oysters from the mosquito control insecticides tested appears low given their estimated and measured

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concentrations in the environment. In the present study, the estimated hazard quotients for resmethrin, permethrin, methoprene, and naled were all far less than one, indicating that use of these compounds for mosquito control, when used as recommended, is unlikely to cause direct mortality to M. mercenaria and C. virginica. Past research has also found that molluscs are the least sensitive among aquatic invertebrates and fishes tested with mosquito control insecticides (Hill 1985; Rand 2002; Solomon et al. 2001). DeLorenzo and Fulton (2012) reported 96-h LC50 values for permethrin of 0.00002 mg/L for mysids and 0.00005 mg/L for grass shrimp (Palaemonetes pugio) compared with a range of 7.65 to [10 mg/L found for the clams and oysters in the present study. Similarly, BoltonWarberg et al. (2007) found adult oysters to be relatively insensitive to OP exposure with a reported 96-h LC50 value of 31.62 mg/L for dichlorvos (breakdown product of naled) compared with 0.062 mg/L for grass shrimp. Zulkosky et al. (2005) detected resmethrin in 50 % of the samples taken from Long Island, New York, within 1 h of spray events at concentrations ranging from 1.7 9 10-6 to 9.8 9 10-4 mg/L. Targeted monitoring in the Florida Keys National Marine Sanctuary after mosquito control spraying detected permethrin in canal surface water at a maximum concentration of 9.4 9 10-3 mg/L; naled was detected once out of 144 total samples at 1.9 9 10-4 mg/ L, and dichlorvos (breakdown product of naled) was detected at concentrations B5.6 9 10-4 mg/L (Pierce et al. 2005). In addition, Bolton-Warberg et al. (2007) did not detect naled in water samples after two spray events in Charleston, South Carolina, but dichlorvos was detected at 2.1 9 10-4 mg/L. Although some measured field values exceed the estimated environmental concentrations used in the hazard assessment, they are still several orders of magnitude lower than the acute toxicity values for mortality or swimming activity determined in this study. Chronic, sublethal effects of the mosquito control insecticides on the growth of M. mercenaria and C. virginica occurred at levels lower than the thresholds for acute mortality, yet these still exceed likely environmental exposure concentrations from mosquito control insecticides (e.g. Scott et al. 2002). The addition of abiotic stressors (hypoxia and hypercapnia) increased juvenile clam mortality with insecticide exposure, but the resmethrin concentration necessary to elicit significant mortality was still lower than insecticide levels likely to occur in the environment (Rand 2002). Expanded pesticide-monitoring programs in marine and estuarine waters are needed to fully evaluate mosquito control chemical exposure to resident aquatic species and to improve the reliability of coastal risk assessments (DeLorenzo and Fulton 2012). Although this assessment suggests that bivalve molluscs are at low risk from mosquito control insecticides, there are

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other factors to consider for a comprehensive analysis of these compounds, e.g. testing the synergized version of mosquito control compounds; testing the compounds together as a mixture; conducting sediment testing with infaunal clams, which could have greater exposure to hydrophobic insecticides after settlement (Spurlock and Lee 2008; Thompson et al. 1999; Weston et al. 2006); and testing other abiotic factors, such as temperature (Talmage and Gobler 2011). Conclusion The results of the present study fill an important gap in our knowledge of insecticide toxicity in bivalve molluscs because threshold concentrations for mortality were mostly unknown in M. mercenaria and C. virginica. Based on the results of this study, current use of mosquito control insecticides poses low risk to native bivalve populations and to coastal shellfish mariculture operations. The study also increased knowledge on sublethal effects resulting from insecticide exposure (decreased swimming activity and growth) and sheds light on the complexity of insecticide toxicity when multiple factors, e.g. chemical class, time of exposure, life stage/species, and effects from abiotic factors, are taken into account. The results of this study specifically showed that levels of low pH caused by increased CO2 and low oxygen currently measured in southeastern estuaries caused significant effects on larval clam survival, which were exacerbated by insecticide exposure. These data will assist environmental resource agencies in managing the use of mosquito control insecticides near sensitive coastal habitats. Acknowledgments This research was funded by a Grant from the Southern Regional Aquaculture Center, in cooperation with the United States Department of Agriculture, Cooperative State Research, Education and Extension Service, to M. E. DeLorenzo and L. D. Coen and the College of Charleston Graduate Program in Marine Biology, Biology Department, and Graduate Student Association. We acknowledge the kind experimental assistance of Paul Pennington, Shannon Whitehead, and John Venturella (NOAA/NOS/CCEHBR) as well as Karen Burnett, Anna Tommerdahl, Rebecca Derex, and Sarah Song (College of Charleston). We thank Mike Fulton, Tina Mikulski, Cheryl Woodley, and Pat Fair for helpful review of the manuscript. The NOAA, National Ocean Service does not approve, recommend, or endorse any proprietary product or material mentioned in this publication. This is Contribution No. 418 of the Grice Marine Laboratory, College of Charleston, Charleston, SC.

References Atlantic States Marine Fisheries Commission (2007) The importance of habitat created by shellfish and shell beds along the Atlantic Coast of the US Prepared by Coen LD and Grizzle R with contributions by Lowery J and Paynter KT Jr. ASMFS, Morehead

Arch Environ Contam Toxicol (2014) 66:450–462 Baker SM, Mann R (1992) Effects of hypoxia and anoxia on larval settlement, juvenile growth, and juvenile survival of the oyster Crassostrea virginica. Biol Bull 182:265–269 Beck MW, Brumbaugh RD, Airoldi L, Carranza A, Coen LD, Crawford C et al (2011) Oyster reefs at risk and recommendations for conservation, restoration, and management. Bioscience 61:107–116 Bolton-Warberg M, Coen LD, Weinstein JE (2007) Acute toxicity and acetylcholinesterase inhibition in grass shrimp (Palaemonetes pugio) and oysters (Crassostrea virginica) exposed to the organophosphate Dichlorvos: Laboratory and field experiments. Arch Environ Contam Toxicol 52:207–216 Bondarenko S, Putt A, Kavanaugh S, Poletika N, Gan J (2006) Time dependence of phase distribution of pyrethroids insecticides in sediment. Environ Toxicol Chem 25:3148–3154 Cai WJ, Wang Y (1998) The chemistry, fluxes, and sources of carbon dioxide in the estuarine waters of the Satilla and Altamaha rivers, Georgia. Limnol Oceanogr 43:657–668 Carriker MR (2001) Embryogenesis and organogenesis of veligers and early juveniles. In: Kraeuter JN, Castagna M (eds) The biology of the hard clam. Elsevier Science BV, Amsterdam, pp 77–112 Chandler GT, Cary TL, Volz DC, Walse SS, Ferry JL, Klosterhaus SL (2004) Fipronil effects on estuarine copepod (Amphiascus tenuiremis) development, fertility, and reproduction: A rapid life-cycle assay in 96-well microplate format. Environ Toxicol Chem 23:117–124 Cochran RE, Burnett LE (1996) Respiratory responses of the salt marsh animals Fundulus heteroclitus, Leiostomus xanthurus, and Palaemonetes pugio to environmental hypoxia and hypercapnia and to the organophosphate insecticide, azinphosmethyl. J Exp Mar Biol Ecol 195:125–144 Coen LD, Heck KL Jr (1991) The interacting effects of siphon nipping and habitat on bivalve (Mercenaria mercenaria (L)) growth in a subtropical seagrass (Halodule wrightii Aschers) meadow. J Exp Mar Biol Ecol 145:1–13 Coen LD, Lukenbach MW (2000) Developing success criteria and goals for evaluating oyster reef restoration: Ecological function or resource exploitation? Ecol Eng 15:323–343 Coen LD, Dumbauld BR, Judge ML (2011) Expanding shellfish aquaculture: A review of the ecological services provided by and impacts of native and cultured bivalves in shellfish-dominated ecosystems. In: Shumway SE (ed) Shellfish aquaculture and the environment. Wiley-Blackwell, New York, pp 239–295 Dame R (1996) Ecology of marine bivalves: an ecosystem approach. CRC Marine Science Series, Boca Raton DeLorenzo ME, Fulton MH (2012) Comparative risk assessment of permethrin, chlorothalonil, and diuron to coastal aquatic species. Mar Pollut Bull 64:1291–1299 DeLorenzo ME, Plante C, Coen LD (2012) Effects of mosquito abatement pesticides on various life stages of commercially important shellfish aquaculture species in the South. Final project report. Southern Regional Aquaculture Center, Stoneville Dickinson GH, Ivanina AV, Matoo OB, Po¨rtner HO, Lannig G, Bock C et al (2012) Interactive effects of salinity and increased CO2 levels on juvenile eastern oysters, Crassostrea virginica. J Exp Biol 215:29–43 Dickinson GH, Matoo OB, Tourek RT, Sokolova IM, Beniash E (2013) Environmental salinity modulates the effects of increased CO2 levels on juvenile hard-shell clams, Mercenaria mercenaria. J Exp Biol 216:2607–2618 Dove MC, Sammut J (2007) Impacts of estuarine acidification on survival and growth of Sydney rock oysters Saccostrea glomerata (Gould 1850). J Shellfish Res 26:519–527 Dumbauld BR, Ruesink JL, Rumrill SS (2009) The ecological role of bivalve shellfish aquaculture in the estuarine environment: a

461 review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290:196–223 Forrest BM, Keeley NB, Hopkins GA, Webb SC, Clement DM (2009) Bivalve aquaculture in estuaries: review and synthesis of oyster cultivation effects. Aquaculture 298:1–15 Grabowski JH, Brumbaugh RD, Conrad RF, Keeler AG, Opaluch JJ, Peterson CH et al (2012) Economic valuation of ecosystem services provided by oyster reefs. Bioscience 62:900–909 Habig C, DiGiulio R (1991) Biochemical characteristics of cholinesterases in aquatic organisms. In: Mineau P (ed) Cholinesteraseinhibiting insecticides: Their impact on wildlife and the environment. Elsevier Science, New York, pp 19–33 Hales S, de Wet N, Maindonald J, Woodward A (2002) Potential effect of population and climate changes on global distribution of dengue fever: an empirical model. Lancet 360:830–834 Hennessey MK, Nigg HN, Habeck DH (1992) Mosquito (Diptera: Culicidae) adulticide drift into wildlife refuges of the Florida Keys. Entomol Soc Am 14:714–720 Henrick CA (2007) Methoprene: biorational control of mosquitoes. J Am Mosq Control Assoc 23:225–239 Hill IR (1985) Effects on non-target organisms in terrestrial and aquatic environments. In: Leahey J (ed) The pyrethroid insecticides. Taylor and Francis, Philadelphia, pp 189–238 Hoguet J, Key PB (2007) Activities of biomarkers in multiple life stages of the model crustacean, Palaemonetes pugio. J Exp Mar Biol Ecol 353:235–244 Holland AF, Sanger DM, Gawle CP, Lerberg SB, Santiago MS, Riekerk GHM et al (2004) Linkages between tidal creek ecosystems and the landscape and demographic attributes of their watersheds. J Exp Mar Biol Ecol 298:151–178 Huba´lek Z, Halouzka J (1999) West Nile fever—a reemerging mosquito-borne viral disease in Europe. Emerg Infect Dis 5:643–650 Jaffe R (1991) Fate of hydrophobic organic pollutants in the aquatic environment: a review. Environ Pollut 69:237–257 Kellogg ML, Cornwell JC, Owens MS, Paynter KT (2013) Denitrification and nutrient assimilation on a restored oyster reef. Mar Ecol Prog Ser 480:1–19 Kemp WM (1989) Estuarine chemistry. In: Day JW, Hall CA, Kemp WM, Yanez-Arancibia A (eds) Estuarine ecology. Wiley, New York, pp 79–145 Kennedy VS, Krantz LB (1982) Comparative gametogenic and spawning patterns of the oyster Crassostrea virginica in central Chesapeake Bay, USA. J Shellfish Res 2:133–140 Key P, DeLorenzo M, Gross K, Chung K, Clum A (2005) Toxicity of the mosquito control pesticide ScourgeÒ to adult and larval grass shrimp (Palaemonetes pugio). J Environ Sci Health B 40:585–594 Kraeuter JN, Castagna M (2000) Predators and predation. In: Kraeuter JN, Castagna M (eds) The biology of the hard clam. Elsevier Science BV, Amsterdam, pp 441–590 Kuffner IB, Andersson AJ, Jokiel PL, Rodgers KS, Mackenzie FT (2008) Decreased abundance of crustose coralline algae due to ocean acidification. Nat Geosci 1:114–117 Kurihara H, Kato S, Ishimatsu A (2007) Effects of increased seawater pCO2 on early development of the oyster Crassostrea gigas. Aquat Biol 1:91–98 Lehotay SJ, Harman-Fetcho JA, McConnell LL (1998) Agricultural pesticide residues in oysters and water from two Chesapeake Bay tributaries. Mar Pollut Bull 37:32–44 Lerberg SB, Holland AF, Sanger DM (2000) Responses of tidal creek macrobenthic communities to the effects of watershed development. Estuaries 23:838–853 Luckenbach MW, Coen LD, Ross PG, Stephen JA (2005) Oyster reef habitat restoration: relationships between oyster abundance and community development based on two studies in Virginia and South Carolina. J Coast Res 40:64–78

123

462 Martens P, Hall L (2000) Malaria on the move: human population movement and malaria transmission. Emerg Infect Dis 6:103–109 Milam CD, Farris JL, Wilhide JD (2000) Evaluating mosquito control pesticides for effect on target and nontarget organisms. Arch Environ Contam Toxicol 39:324–328 Miller TA, Salgado VL (1985) The mode of action of pyrethroids on insects. In: Leahy JP (ed) The pyrethroid insecticides. Taylor and Francis, London, pp 43–97 Miller AW, Reynolds AC, Sobrino C, Riedel GF (2009) Shellfish face uncertain future in a high CO2 world: Influence of acidification on oyster larvae calcification and growth in estuaries. PLoS One 4:e5661 National Marine Fisheries Service (2011) Fisheries of the United States 2010. Office of Science and Technology, Silver Spring Naylor RL, Goldburg RJ, Primavera JH, Kautsky N, Beveridge MCM, Clay J et al (2000) Effect of aquaculture on world fish supplies. Nature 405:1017–1024 Newell RIE (2004) Ecosystem influences of natural and cultivated populations of suspension-feeding bivalve mollusks: a review. J Shellfish Res 23:51–61 Norberg-King TJ (1993) A linear interpolation method for sublethal toxicity: the inhibition concentration (ICP) approach. Technical report 03-93. United States Environmental Protection Agency, Duluth North EW, Schlag Z, Hood RR, Li M, Zhong L, Gross T et al (2008) Vertical swimming behavior influences the dispersal of simulated oyster larvae in a coupled particle-tracking and hydrodynamic model of Chesapeake Bay. Mar Ecol Prog Ser 359:99–115 Peterson CH, Grabowski JH, Powers SP (2003) Estimated enhancement of fish production resulting from restoring oyster reef habitat: quantitative valuation. Mar Ecol Prog Ser 264:249–264 Pierce RH, Henry MS, Blum TC, Mueller EM (2005) Aerial and tidal transport of mosquito control pesticides into the Florida Keys National Marine Sanctuary. Rev Biol Trop 53:117–125 Rand GM (2002) Hazard assessment of resmethrin: effects and fate in aquatic systems. Ecotoxicology 11:101–111 Reichenberger S, Bach M, Stikschak A, Frede HG (2007) Mitigation strategies to decrease pesticide inputs into ground- and surface water and their effectiveness: a review. Sci Total Environ 384:1–35 Roop T, Greenberg MJ (1967) Acetylcholinesterase activity in Crassostrea virginica and Mercenaria mercenaria. Am Zool 7:737–738 Scott GI, Fulton MH, Wirth EF, Chandler GT, Key PB, Daugomah JW et al (2002) Toxicological studies in tropical ecosystems: an ecotoxicological risk assessment of pesticide runoff in south Florida estuarine ecosystems. J Agric Food Chem 50:4400–4408 Shumway SE (1996) Natural environmental factors. In: Kennedy VS, Newell RIE, Eble AF (eds) The eastern oyster Crassostrea virginica. Maryland Sea Grant, College Park, pp 467–513 Soderlund DM, Bloomquist JR (1989) Neurotoxic actions of pyrethroid insecticides. Annu Rev Entomol 34:77–96

123

Arch Environ Contam Toxicol (2014) 66:450–462 Solomon KR, Giddings JM, Maund SJ (2001) Probabilistic risk assessment of cotton pyrethroids: I. Distributional analyses of laboratory aquatic toxicity data. Environ Toxicol Chem 20:652–659 Spurlock F, Lee M (2008) Synthetic pyrethroid use patterns, properties, and environmental effects. In: ACS symposium series. American Chemical Society, Washington. pp 3–25 Stunz GW, Minello TJ, Rozas LP (2010) Relative value of oyster reef as habitat for estuarine nekton in Galveston Bay, Texas. Mar Ecol Prog Ser 406:147–159 Suter GW II (1995) Introduction to ecological risk assessment for aquatic toxic effects. In: Rand GM (ed) Fundamentals of aquatic toxicology, 2nd edn. Taylor and Francis, Washington, pp 803–816 Talmage SC, Gobler CJ (2011) Effects of increased temperature and carbon dioxide on the growth and survival of larvae and juveniles of three species of Northwest Atlantic bivalves. PLoS One 6:e26941 Tamburri MN, Zimmer-Faust RK, Tamplin ML (1992) Natural sources and properties of chemical inducers mediating settlement of oyster larvae: a re-examination. Biol Bull 183:327–338 Thompson B, Anderson B, Hunt J, Taberski K, Phillips B (1999) Relationships between sediment contamination and toxicity in San Francisco Bay. Mar Environ Res 48:285–309 Tomanek L, Zuzow MJ, Ivanina AV, Beniash E, Sokolova IM (2011) Proteomic response to increased PCO2 level in eastern oysters, Crassostrea virginica: Evidence for oxidative stress. J Exp Biol 214:1836–1844 Waldbusser GG, Steenson RA, Green MA (2011) Oyster shell dissolution rates in estuarine waters: effects of pH and shell legacy. J Shellfish Res 30:659–669 Watson S, Southgate PC, Tyler PA, Peck LS (2009) Early larval development of the Sydney rock oyster Saccostrea glomerata under near-future predictions of C02-driven ocean acidification. J Shellfish Res 28:431–437 Weston DP, Amweg EL, Mekebri A, Ogle RS, Lydy MJ (2006) Aquatic effects of aerial spraying for mosquito control over an urban area. Environ Sci Technol 40:5817–5822 Wheeler MW, Park RM, Bailer AJ (2006) Comparing median lethal concentration values using confidence interval overlap or ratio tests. Environ Toxicol Chem 25:1441–1444 Wilson JD (2002) Productivity, fisheries, and aquaculture in temperate estuaries. Est Coast Shelf Sci 55:953–967 Zar JH (1999) Biostatistical analysis, 4th edn. Prentice-Hall, Upper Saddle River Zulkosky AM, Ruggieri JP, Terracciano SA, Brownawell BJ, McElory AE (2005) Acute toxicity of resmethrin, malathion, and methoprene to larval and juvenile American lobsters (Homarus americanus) and analysis of pesticide levels in surface waters after ScourgeÒ, AnvilÒ, and AltosidÒ application. J Shellfish Res 24:795–804