Printer Friendly

Effects of mosquito control pesticides on competent queen conch (Strombus gigas) larvae.


Mosquitoes pose a health threat and are a nuisance to humans in the Florida Keys. As such, mosquito control pesticides are regularly employed, especially during the rainy summer season, when mosquitoes are more prevalent. Two mosquito adulticides used by the Florida Keys Mosquito Control District are naled and permethrin (brand names Dibrom and Biomist 30-30, respectively). Naled is an organophosphate pesticide designed to target the nervous system of insects by inhibiting the enzyme acetylcholinesterase, which is involved in the transmission of nerve impulses (Cox, 2002). Naled's breakdown product, dichlorvos, works similarly (Cox, 2002). Permethrin is a synthetic pyrethroid whose mode of action also targets the nervous system by acting as a sodium-channel blocker (Cox, 1998). Both are sprayed over land as an ultra-low-volume mist, naled from aircraft and permethrin from a truck-mounted mister. Research has shown that aerial drift and runoff can carry these pesticides into non-target areas and nearshore waters in the Keys (Hennessey et al., 1992; Pierce et al, 2005).

The queen conch (Strombus gigas Linnaeus, 1758) is a marine gastropod that inhabits the tropical western Atlantic including the Keys. It once supported significant commercial and recreational fisheries in the Keys, but because of a decline in the stock, harvest has been prohibited since 1985. The population has been slow to recover, and current levels still cannot support exploitation (Glazer and Delgado, 2003; Florida Fish and Wildlife Conservation Commission, un-publ. data). The prolonged recovery has been attributed, in part, to limited larval recruitment (Stoner et al., 1996a, 1997; Glazer and Delgado, 2003; Delgado et al., 2008). Queen conch in the Keys breed during the spring and summer (Delgado et al., 2004); consequently, their larvae are most abundant when pesticide use is at its peak. Furthermore, queen conch larvae exhibit positive phototaxis and are thus associated with surface layers (Barile et al., 1994; Stoner and Davis, 1997), where many contaminants, including pesticides, accumulate (Rumbold and Snedaker, 1997).

A great deal of work has been done describing the adverse effects--lethal and sublethal--of naled, dichlorvos, and permethrin on numerous non-target marine invertebrates, including molluscs. For example, naled exposure resulted in delayed development, deformities, or death in queen conch embryos (McIntyre et al., 2006). Dichlorvos reduced the feeding rate and altered behavior in mussels (Donkin et al., 1997) and inhibited acetylcholinesterase activity in oysters (Bolton-Warberg et al., 2007). Permethrin exposure caused abnormal development in oyster larvae (Cox, 1998) and queen conch embryos (Mcintyre et al., 2006).

These observations suggest that the pesticides used for mosquito eradication in the Keys have the potential to negatively affect queen conch larvae. However, none of these studies examined the effects of these pesticides on metamorphic success. Metamorphosis is a seminal event characterized by a suite of morphological and physiological changes that occur as larvae move from a planktonic to a benthic way of life, and inopportune exposure to pesticides may have a deleterious impact (sensu Rodriguez et al., 1993). To examine this issue, we exposed competent (i.e., capable of undergoing metamorphosis) queen conch larvae to naled and permethrin to determine the pesticides' lethal and sublethal effects on this crucial early life-history stage.

Materials and Methods

Test animals

Queen conch larvae were cultured using the techniques described by Davis (1994a) and briefly outlined here. Egg masses were obtained from wild conch located on the back reef of the Keys reef tract. The egg masses were shipped to Harbor Branch Oceanographic Institute at Florida Atlantic University (HBOI-FAU) for incubation and larval rearing. Larvae were raised in 700-1 fiberglass tanks for 21 days. At this point, a number of morphological changes occur that signal metamorphic competency. These include eye migration toward the end of the eyestalks and a change in the pigment on the foot from orange to dark green (Davis et al., 1990; Davis, 1994a).

The best natural metamorphic inducer for queen conch is an extract of the red alga Laurencia potei (Lamouroux) Howe, 1918 (Davis et al., 1990; Davis, 1994a). On Day 18 of rearing, a dosage test was performed on a subsample of larvae to determine the lowest concentration of L. potei extract required to induce metamorphosis in more than 50% of larvae after a 3-h exposure. The extract was prepared, and the dosage test was performed using the procedures described by Davis et al (1990). A successful metamorphosis is recognized when the velar lobes have been lost and the animal has begun crawling on its foot (Davis et al., 1990; Davis, 1994a). Our research partners at HBOI-FAU determined that 1 ml of extract per 100 ml of seawater was suitable for inducing metamorphosis (Amber Garr, pers. comm.). Three days after this dosage was determined (Day 21), the remaining larvae were transported to our laboratory in the Keys and used in the exposure experiments later in the day of arrival.

Pesticide exposure procedures

Exposures were conducted as static tests performed in glass petri dishes (150-ml capacity). Temperature in the dishes was maintained at 28 [degrees]C in a temperature-controlled room; the room also had a 12-h light-dark cycle. We used Instant Ocean synthetic seawater at a salinity of 35%0 for the exposure solutions. Prior to use, the synthetic seawater was filtered through a 5-[micro]m charcoal filter, a 5-[micro]m cellulose filter, and a 1-[micro]m cellulose filter to remove impurities and particulates.

Competent queen conch larvae were exposed to treatments containing either naled or permethrin. Each experiment consisted of a control of synthetic seawater and five pesticide concentrations, for a total of six treatments. Treatments had three replicates; each replicate had 10 larvae each. Larvae were randomly assigned to each treatment. Test concentrations were 0 ng/ml (control), 1.75 ng/ml, 3.75 ng/ml, 7.5 ng/ml, 15 ng/ml, and 30 ng/ml. These values were environmentally relevant as they encompassed the range that Pierce et al. (2005) found in nearshore waters of the Keys as well as the spray concentration (30 ng/ml) used by Florida Keys Mosquito Control. Since pesticide exposures in the natural environment tend to be episodic and of short duration (Forbes and Cold, 2005; Oros and Werner, 2005), we endeavored to emulate environmental conditions and used a 12-h dosing period.

We examined the effect of the pesticides on mortality and metamorphic success. After the 12-h dosing period, we recorded larval mortality and metamorphosis and then used a pipette to transfer the larvae into new petri dishes containing pesticide-free synthetic seawater. At this time, the L. potei extract was added to all treatments, including the control. Larvae were exposed to the extract for 3 h. Larval mortality and metamorphosis were recorded again at 15 h (i.e., 3 h after adding the L. potei extract) and at 48 h. We evaluated mortality and metamorphic success among the treatments using a one-way ANOVA. We also calculated the lowest observed effective concentration (LOEC), defined as the lowest pesticide concentration that was significantly different from the control as determined by the post hoc, multiple-comparison Dunnett test (one-tailed). Statistical tests were run on SPSS 11.0 for Windows. Results were considered significant at P < 0.05.

Pesticide concentration verification

Treatment solutions were analyzed to verify pesticide concentrations and, in the case of naled, the concentration of its degradation product, dichlorvos. The methods briefly described below were based on the Florida Department of Environmental Protection's standard operating procedure for extracting pesticides from water (Reddy et al., 2005). Pesticide concentrations were determined by gas chromatography using a Varian 3800 gas chromatograph with dual-column electron-capture detectors (GC-ECD) and a Turbochrome chromatography data-processing system. Samples of each exposure solution were collected at time 0 and at 12 h and placed in clean, dichloromethane-rinsed amber bottles fitted with a Teflon-lined cap. Dichloromethane was added to each sample as a preservative. The samples were then stored at 4 [degrees]C for subsequent extraction at Mote Marine Laboratory.

Samples were allowed to warm to room temperature and extracted three times via liquid-liquid extraction (in sepa-ratory funnels) with pesticide-grade dichloromethane. The samples were then reduced in volume using the Rapid-Vap system under a stream of high-purity nitrogen gas; the solvent was replaced with pesticide-grade hexane and brought to a volume of 1 ml. The samples were then injected into the dual-column GC-ECD using a Varian 8200 auto-sampler. Each sample was analyzed a minimum of two times to check instrument variability. Blanks were run between samples to ensure that there was no sample carryover. Calibration curves and relative response factors were calculated; in addition, pesticide standards were run to monitor instrument sensitivity. The minimum detectable levels for naled, dichlorvos, and permethrin were 0.010 ng/ml, 0.006 ng/ml, and 0.005 ng/ml, respectively.


Pesticide concentration verification

Nominal pesticide concentrations for the exposure experiments ranged from no detection (ND) in the control to 30 ng/ml for both naled and permethrin. However, actual concentrations as determined analytically tended to be lower. Nominal and actual concentrations are listed in Table 1.

In the naled exposures, actual concentrations at time 0 ranged from ND in the control to 28.28 ng/ml in the 30 ng/ml treatment. Concurrently, levels of dichlorvos (naled's breakdown product) ranged from ND in the control to 6.30 ng/ml in the 30 ng/ml naled treatment. Concentrations of both naled and dichlorvos had decreased by the 12-h sampling period. In general, naled decreased by an order of magnitude, with no detectable naled in the control and lowest treatment concentration. There was no detectable dichlorvos in the control or the two lowest treatment concentrations at 12 h; dichlorvos was more persistent at the higher treatment concentrations.

Actual permethrin concentrations were much lower than our nominal targets (see Table 1 for concentrations). These concentrations ranged from ND in the control to only 5.25 ng/ml in the 30 ng/ml treatment at time 0 (Table 1). Although the highest actual concentrations were not as high as that of the Florida Keys Mosquito Control spray concentration (30 ng/m1), they are similar to the highest values of permethrin found by Pierce et al. (2005) in Keys nearshore waters (i.e., 5.1 to 9.4 ng/ml). In general, the concentration of permethrin at the 12-h sampling period had decreased by an order of magnitude, with no detectable levels of permethrin in the control and two lowest treatment concentrations (Table 1).

Table 1

Nominal and actual pesticide concentrations (ng/ml)
during the 12-hour exposure period

                             Actual Conc

                    Naled    Dichlorvos   Permethrin

Conc.          0 h   12 h    0 h   12 h    0 h  12 h

0 (control)   0.00   0.00   0.00   0.00   0.00  0.00

1.75          0.04   0.00   0.03   0.00   0.28  0.00

3.73          1.05   0.17   0.06   0.00   0.84  0.00

7.5           4.09   0.89   1.46   0.38   1.55  0.33

15            9.77   3.86   3.80   4.13   4.68  0.45

30           26.28   6.59   6.30   5.70   5.25  1.55

Pesticide exposure: naled

The competent queen conch larvae used in this experiment experienced very little to no mortality. No mortality was observed in any of the treatments after the 12-h exposure to naled. Three h after introducing the L. potei extract (i.e., 15 h after initial pesticide exposure). no mortality was observed; however, some larvae appeared to be in poor condition. At 48 h. four of the six treatments experienced 3% to 7% mortality; none of the individuals in the two highest treatment concentrations suffered any mortality (Fig. 1). However, these differences were not statistically significant (ANOVA: [F.sub.[5, 12]]. 121 = 0.686, P = 0.643) (Fig. 1).

No metamorphosis was observed in any of the treatments after the 12-h exposure to naled. Metamorphosis data at 15 h (i.e., 3 h after introducing the L. potei extract) and at 48 h were identical; thus, only the 48-h data are presented. There was a significant difference among the treatments in the proportion of larvae that metamorphosed successfully (ANOVA: [F.sub.[5, 12]] = 8.028, P = 0.002) (Fig. 1). Metamor-phic success increased with increasing naled concentration (Fig. 1); a linear regression of metamorphosis success on actual naled concentration showed a significant positive relationship ([[gamma].sup.2] = 0.470, P = 0.002). The LOEC for naled (i.e., the lowest actual treatment concentration that was significantly greater than the control) was 26.28 ng/ml (P = 0.025); the corresponding dichlorvos value was 6.30 ng/ml (Table 1). The 26.28 ng/ml treatment had a metamorphic success rate of about 87% compared with 57% in the control (Fig. 1).

Pesticide exposure: permethrin

The results from the permethrin exposures were similar to those for naled. No mortality was observed in any of the treatments after the 12-h exposure to permethrin. No mortality was observed 3 h after introducing the L. potei extract (i.e., 15 h after initial pesticide exposure). At 48 h, nominal treatments 7.5 ng/ml (1.55 ng/ml actual) and 30 ng/ml (5.25 ng/ml actual) suffered 3% mortality; the other treatments did not experience any mortality (Fig. 2). However, these differences were not statistically significant (ANOVA: [F.sub.[5, 12]] = 0.800, P = 0.571) (Fig. 2).

After the 12-h exposure to permethrin, a single larva underwent metamorphosis in the nominal 15 ng/ml treatment (4.68 ng/ml actual); no metamorphosis was observed in any of the other treatments after 12 h of exposure to permethrin. The metamorphosis data at 15 h (i.e., 3 h after introducing the L. potei extract) and at 48 h were very similar; thus, only the 48-h data are presented. There was a significant difference in metamorphic success among the treatments (ANOVA: [F.sub.[5, 12]] = 5.738, P = 0.006) (Fig. 2). As for naled, metamorphic success increased with increasing permethrin concentration (Fig. 2); a linear regression of metamorphic success on actual permethrin concentration showed a significant positive relationship ([[gamma].sup.2] = 0.342, P = 0.011). The LOEC for permethrin was 4.68 lig/ml (P = 0.008); the 4.68 ng/ml treatment had a metamorphic success rate of about 85% compared to 43% in the control (Fig. 2).


An extensive body of literature details the toxicity of naled, dichlorvos, and permethrin to a variety of non-target marine invertebrates, including molluscs. For instance, permethrin was found to cause 100% mortality in newly hatched queen conch larvae after 48 h (Mcintyre et al., 2006). However, our results indicate that these mosquito adulticides do not seem to be especially toxic to competent queen conch larvae at the environmentally relevant concentrations to which the larvae were exposed. The very low mortality (Figs. 1 and 2) is probably due to the 12-h exposure period used in this study (which is a more realistic exposure scenario than the standard 96-h. static-toxicity test) and that the pesticides of interest were designed to kill arthropods, not molluscs. For example, Bolton-Warberg et al. (2007) found that, based on L[C.sub.50] values, grass shrimp were more sensitive than oysters to dichlorvos. Similarly, Clark et al. (1989) and Oros and Werner (2005) reported that marine crustaceans were more sensitive than oysters to permethrin.

In addition to having no directly lethal effects, naled, dichlorvos, and permethrin did not induce metamorphosis, as only a single larva in one of the permethrin treatments metamorphosed after the 12-h exposure period. Therefore, the pesticides did not act as a false metamorphic cue. Nor did the pesticides interfere with metamorphosis. In fact, after the introduction of the L. potei extract, metamorphic success was higher in the larvae that had been exposed to the higher pesticide concentrations (Figs. 1 and 2).

Mollusc= metamorphosis is governed by two independent chemosensory pathways, a morphogenetic pathway that induces metamorphosis when stimulated and a regulatory pathway that amplifies sensitivity to metamorphic cues (Baxter and Morse, 1987; Morse, 1993; Boettcher and Tar-gett, 1998). Our results suggest that the pesticides activated the regulatory pathway and enhanced the queen conch larvae's response to the natural metamorphic inducer (i.e., the L. potei extract). The regulatory pathway has been hypothesized to play a role in the identification of high-quality habitat for metamorphosis, and thus, favorable conditions for the postlarval organism (Baxter and Morse, 1987; Morse, 1993).

The sublethal effects of pesticide exposure may be lethal in the long run by reducing an individual's fitness, ultimately leading to mortality and population declines (Oros and Werner, 2005), as has been shown in California with amphibians (Spading et at., 2001; Davidson, 2004). Aerial drift and runoff can carry naled, dichlorvos, and permethrin into Keys nearshore waters (Pierce et al., 2005). Our results suggest that these pesticides may sensitize queen conch larvae to metamorphic cues. Queen conch larvae are known to metamorphose in response to habitat and trophic cues, which improves the probability that larvae will recruit to areas that will provide suitable forage for postlarval juveniles (Davis and Stoner, 1994; Boettcher and Targett, 1996; Stoner et at., 1996b). In this vein, Kowalik et al. (2006) found that newly settled juvenile queen conch from low-quality habitat treatments were not as robust (defined as actively crawling and searching for food with the proboscis) as those in high-quality treatments. Therefore, if queen conch larvae metamorphose in suboptimal habitat due to increased sensitivity to metamorphic cues from pesticide exposure, recruit survival will most Rely decrease. Consequently, mosquito adulticides may have a negative impact on the recovery of nearshore queen conch populations by adversely affecting recruitment in a population that is already recruitment limited (Stoner et al., 1996a, 1997; Glazer and Delgado 2003; Delgado et al, 2008) due to small spawning aggregations (Glazer and Delgado, 2003; Delgado et at., 2008; Florida Fish and Wildlife Conservation Commission, unpubl. data).

Considerable concern has been expressed regarding the extrapolation of laboratory-derived toxicity data to field situations; it has been argued that laboratory tests can be applied to field conditions only when the exposure regime used is comparable to that found in the natural environment (Forbes and Cold, 2005; Oros and Werner, 2005). Our exposure regime was designed to mimic environmental conditions by exposing competent queen conch larvae to a 12-h dosing period with environmentally relevant pesticide concentrations. We attempted to bracket the range that Pierce et al. (2005) found in nearshore waters of the Keys with the spray concentration used by Florida Keys Mosquito Control (30 ng/ml). The actual concentrations for the naled exposures were quite close to their nominal values (Table 1). However, the insolubility of pyrethroids (Oros and Werner, 2005) prevented us from reaching our targeted permethrin concentrations (Table 1). Nevertheless, the actual values fell within the concentrations found by Pierce et al. (2005); therefore, our results are pertinent to potential field exposures in the Keys.

Our results also highlight the importance of choosing germane endpoints, as the pesticides did not have a directly lethal effect on competent queen conch larvae, but did have the sublethal post-exposure effect of increasing metamorphic success with increasing pesticide concentration (Figs. 1 and 2). Metamorphic success at our LOECs was about twice that in the controls. A comparison between pesticide concentrations found by Pierce et al. (2005) and our LOECs shows that naled (0.19 ng/ml) and dichlorvos (0.60 ng/ml) in Keys nearshore waters were well below our LOECs (26.28 ng/ml and 6.30 ng/ml, respectively). However, Pierce et al. (2005) found permethrin concentrations as high as 9.40 ng/ml in surface waters, almost double our LOEC of 4.68 ng/ml. On the basis of these comparisons, permethrin may pose a greater risk to competent queen conch larvae in Keys nearshore waters than naled or dichlorvos.

Clearly, more research is needed on the effects that inopportune pesticide exposure may have on queen conch larvae. For example, late-stage precompetent larvae may undergo a forced early metamorphosis if sensitized to metamorphic cues by pesticide exposure. Previous research has shown that juveniles derived from precompetent larvae were lethargic and emaciated, reducing the likelihood of juvenile success (Davis, 1994b). In addition, there may be synergistic effects if larvae are exposed to both pesticides at the same time or if pesticides are introduced in concurrence with metamorphic cues. Further studies are also warranted on the long-term impacts of pesticide exposure. Questions remain as to whether newly metamorphosed postlarvae can develop normally, reproduce successfully, and produce viable young after pesticide exposure or whether long-term sublethal effects, such as endocrine disruption (sensu Delgado et al., 2004; De Guise et al., 2005; Oros and Werner, 2005; Walker er al., 2005), may be exacerbating the species' protracted recovery in the Florida Keys.


This paper is funded, in part, by a grant/cooperative agreement from the South Florida Water Management District (SFWMD Agreement No. OT050676). The views expressed herein are those of the authors and do not necessarily reflect the views of the SFWMD or any of its sub-agencies. Additional funding was provided by the Florida Fish and Wildlife Conservation Commission. Phil Mercurio, Maria Baron. and Shoshana Whiner assisted with the experiments. We thank Amber Garr at HBOI-FAU for culturing the queen conch larvae. Steve Geiger, Ted Lange, Bland Crowder, and two anonymous reviewers provided insightful comments on earlier versions of the manuscript.

Received 24 June 2013; accepted 16 October 2013.

* To whom correspondence should be addressed. E-mail: gabriel.

Abbreviations: LOEC, lowest observed effective concentration.

Literature Cited

Bark, P. J., A. W. Stoner, and C. M. Young. 1994. Phototaxis and vertical migration of the queen conch (Strombus gigas linne) veliger larvae. J. Exp. Mar Biol. Ecol. 183: 147-162,

Baxter, G., and D. E. Morse. 1987. G protein and diacylglycerol regulate metamorphosis of planktonic molluscan larvae. Proc. Natl. Acad. Sci. USA 84: 1867-1870.

Boettcher, A. A., and N. M. Targett. 1996. Induction of metamorphosis in queen conch. Strombus gigas Linnaeus, larvae by cues associated with red algae from their nursery grounds. J. Exp. Mar. Biol. Earl. 196: 29-52.

Boettcher, A. A., and N. M. Targett. 1998. Role of chemical inducers in larval metamorphosis of queen conch, Strombus gigas Linnaeus: relationship to other marine invertebrate systems. Biol. Bull. 194: 132-142.

Bolton-Warberg, M., L. D. Coen, and J. E. Weinstein. 2007. Acute toxicity and acetylcholinesterase inhibition in grass shrimp (Palaetnon-etes pugio) and oysters (Crassostrea virginica) exposed to the organophosphate dichlorvos: laboratory and field studies. Arch. Environ. Contam. Toxicol. 52: 207-216.

Clark, J. R., L. R. Goodman, P. W. Borthwick, J. M. Patrick, G. M. Cripe, P. M. Moody, J. C. Moore, and E. M. Lores. 1989. Toxicity of pyrethroids to marine invertebrates and fish: a literature review and test results with sediment-sorbed chemicals. Environ. Toxicol. Chem. 8: 393-401.

Cox, C. 1998. Permethrin insecticide factsheet. Pestic. Reform 18: 14-20.

Cox, C. 2002. Naled (Dibrom) insecticide factsheet. J. Pestic. Reform 22: 16-21.

Davidson, C. 2004. Declining downwind: amphibian population declines in California and historical pesticide use. Ecol. Appl. 14: 18921902.

Davis, M. 1994a. Mariculture techniques for queen conch (Strombus gigas L.): egg mass to juvenile stage. Pp. 231-252 in Strombus gigas: Queen Conch Biology, Fisheries, and Mariculture, R. S. Appeldoorn and B. Rodriguez, eds. Fundacion Cientifica Los Rogues. Caracas, Venezuela.

Davis, M. 1994b. Short-term competence in larvae of queen conch Strombus gigas: shifts in behavior, morphology, and metamorphic response. Mar. Eral. Prog. Ser. 104: 101-108.

Davis, M., and A. W. Stoner. 1994. Trophic cues induce metamorphosis of queen conch larvae (Strombus gigas Linnaeus). J. Exp. Mar. Biol. Ecol. 180: 83-102.

Davis, M., W. D. Heyman, W. Harvey, and C. A. Withstandley. 1990. A comparison of two inducers, KCJ and Laurencia extracts, and techniques for the commercial scale induction of metamorphosis in queen conch Strombus gigas Linnaeus, 1758 larvae. J. Shellfish Res. 9: 67-73.

De Guise, S., J. Maratea, E. S. Chang, and C. Perkins. 2005. Res-methrin immunotoxicity and endocrine disrupting effects in the American lobster (Homarus americanus) upon experimental exposure. J. Shellfish Res. 24: 781-786.

Delgado, G. A., C. T. Bartels, R. A. Glazer, N. J. Brown-Peterson and K. J. McCarthy. 2004. Translocation as a strategy to rehabilitate the queen conch (Strombus gigas) population in the Florida Keys. Fish. Bull. 102: 278-288.

Delgado, G. A., R. A. Glazer, D. Hawtof, D. Aldana Aranda, L. A. Rodriguez-Gil, and A. de Jestis-Navarrete. 2008. Do queen conch (Strombus gigas) larvae recruiting to the Florida Keys originate from upstream sources? Evidence from plankton and drifter studies. Pp. 29-41 in Caribbean Connectivity: Implications for Marine Protected Area Management. R. Grober-Dunsmore and B. D. Keller. eds. Marine Sanctuaries Conservation Series ONMS-08-07, National Oceanic and Atmospheric Administration, Silver Spring. MD.

Donkin, P., J. Widdows, S. V. Evans, F. J. Staff, and T. Yan. 1997.

Effect of neurotoxic pesticides on the feeding rate of marine mussels (Mytilus edulis). Pest. Manag. Sci. 49: 196-209.

Forbes, V. E., and A. Cold. 2005. Effects of the pyrethroid esfenvalerate on life-cycle traits and population dynamics of Chironomus riparius--importance of exposure scenario. Environ. Toxieol. Chem. 24: 78-86.

Glazer, R. A., and G. A. Delgado. 2003. Towards a holistic strategy to managing Florida's queen conch (Strornbus gigas) population. Pp. 73-80 in El Caracol Strombus gigas: conocimiento integral para su manejo sustentable en el Caribe, D. Aldana Aranda, ed. CYTED. Programa lberoamericano de Ciencia y Technologia para el Desarrollo, Yucatan, Mexico.

Hennessey, M. K., H. N. Nigg, and D. H. Habeck. 1992. Mosquito (Diptera: Culicidae) adulticide drift into wildlife refuges of the Florida Keys. Environ. Entomol. 21: 714-721.

Kowalik, G., M. Davis, A. Shawl, R. A. Glazer, G. A. Delgado, and C. Evans. 2006. Metamorphic response of queen conch (Strornbus gigas) larvae exposed to sediment and water from nearshore and offshore sites in the Florida Keys. Proc. Gulf Caribb. Fish. Inst. 57: 717-729.

Mcintyre, M., R. A. Glazer, and G. A. Delgado. 2006. The effects of the pesticides Biomist 30/30 and Dibrom on queen conch (Strombus gigas) embryos and larvae: a pilot study. Proc. Gulf Caribb. Fish. Inst. 57: 731-742.

Morse, D. E. 1993. Signaling in planktonic larvae. Nature 363: 406.

Oros, D. R., and I. Werner. 2005. Pyrethroid insecticides: an analysis of use patterns, distributions, potential toxicity and fate in the Sacramento-San Joaquin Delta and Central Valley. White Paper for the Interagency Ecological Program. SFEI Contribution 415. San Francisco Estuary Institute, Oakland, CA.

Pierce, R. H., M. S. Henry, T. C. Blum, and E. M. Mueller. 2005. Aerial and tidal transport of mosquito control pesticides into the Florida Keys National Marine Sanctuary. Rev. Biol. Trop. 53: 117-125.

Reddy, S. M., M. Clague, and M. A. Ghaffari. 2005. Extraction of organophosphorus, organonitrogen and organohalide pesticides, PCBs and PAHs from water, wastes, sediment, PUFs, and filters for subsequent analysis by GC-ECD/NPD/FPD/FID/MS. Standard Operating Procedure GC-002-2.7. Florida Department of Environmental Protection, Tallahassee, FL.

Rodriguez, S. R., F. P. Ojeda, and N. C. Inestrosa. 1993. Settlement of benthic marine invertebrates. Mar. Ecol. Prog. Ser. 97: 193-207.

Rumbold, D. G., and S. C. Snedaker. 1997. Evaluation of bioassays to monitor surface microlayer toxicity in tropical marine waters. Arch. Environ. Contam. Toxicol. 32: 135-140.

Sparling, D. W., G. M. Fellers, and L. L. McConnell. 2001. Pesticides and amphibian population declines in California. USA. Environ. Toxicol. Chem. 20: 1591-1595.

Stoner, A. W., and M. Davis. 1997. Abundance and distribution of queen conch veligers (Strombus gigas Linne) in the central Bahamas. II. Vertical patterns in nearshore and deep-water habitats. J. Shellfish Res. 16: 19-29.

Stoner, A. W., R. A. Glazer, and P. J. Barile. 1996a. Larval supply to queen conch nurseries: relationships with recruitment process and population size in Florida and the Bahamas. J. Shellfish Res. 15: 407-420.

Stoner, A. W., M. Ray, R. A. Glazer, and K. J. McCarthy. 1996b. Metamorphic responses to natural substrata in a gastropod larva: decisions related to postlarval growth and habitat preference. J. Exp. Mar. Biol. Ecol. 205: 229-243.

Stoner, A. W., N. Mehta, and T. N. Lee. 1997. Recruitment of Strom-bus veligers to the Florida Keys reef tract: relation to hydrographic events. J. Shellfish Res. 16: 1-6.

Walker, A. N., P. Bush, T. Wilson, E. S. Chang, T. Miller, and M. N. Horst. 2005. Metabolic effects of acute exposure to methoprene in the American lobster. Homarus americanus. J. Shellfish Res. 24: 787-794.


(1) Florida Fish and Wildlife Conservation Commission, Fish and Wildlife Research Institute, 2796 Overseas Highway, Suite 119, Marathon, Florida 33050; and (2) Mote Marine Laboratory, 1600 Ken Thompson Parkway, Sarasota, Florida 34236
COPYRIGHT 2013 University of Chicago Press
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2013 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:Delgado, Gabriel A.; Glazer, Robert A.; Wetzel, Dana
Publication:The Biological Bulletin
Article Type:Report
Geographic Code:1USA
Date:Oct 1, 2013
Previous Article:Quantitative ethogram of male reproductive behavior in the South European toothcarp aphanius fasciatus.
Next Article:Are you positive? electric dipole polarity discrimination in the yellow stingray, Urobatis jamaicensis.

Terms of use | Privacy policy | Copyright © 2019 Farlex, Inc. | Feedback | For webmasters