Determining the Appropriate Duration of Toxicity Tests with Glochidia of Native Freshwater

Final Completion Report for the period August 1, 2010 – July 31, 2012

Submitted to Edward Hammer U.S. Environmental Protection Agency 77 West Jackson Blvd. Chicago, IL 60604-3590

by

Robert B. Bringolf, Ph.D. The University of Georgia Warnell School of Forestry & Natural Resources 180 East Green St., Athens, GA 30602-2152 Telephone: 706-542-1477 E-mail: [email protected]

M. Christopher Barnhart, Ph.D. Biology Department Missouri State University 901 South National Ave. Springfield, MO 65897 Telephone: 417-836-5166 Email: [email protected]

and

W. Gregory Cope, Ph.D. North Carolina State University Department of Environmental & Molecular Toxicology Box 7633, Raleigh, NC 27695-7633 Telephone: 919-515-5296 E-mail: [email protected]

Submitted May 8, 2013

1

Executive Summary

The overall goal of this project was to provide information for determining the appropriate duration of toxicity tests with the larval stage (glochidia) of freshwater mussels belonging to the family . The project was developed to provide insight into the ecological relevance of using the glochidia closing response to salt as an endpoint for toxicity tests and as a measure of condition in determining the appropriate duration of standardized toxicity tests with glochidia. Specific objectives were to determine: 1) if the duration of glochidia viability, defined as the ability to close in response to salt (NaCl) solution, is equivalent to the duration of infectivity- the ability to attach to a host fish and metamorphose successfully into the juvenile stage; 2) if the duration of infectivity of glochidia deposited in natural stream sediment is similar to the duration observed in water-only exposures; and 3) the probability that glochidia deposited onto the sediment surface will infect host fish, using representative -host species pairs.

During the first phase of the project, we tested glochidia viability and infectivity of six mussel species that differ in their host-infection strategies. Three of these species use mantle lures and host extraction (Lampsilis siliquoidea, Lampsilis cardium, Villosa lienosa), two species broadcast glochidia into the water column ( and Amblema plicata), and

one species releases glochidia in conglutinates (Ptychobranchus occidentalis). Our results

indicate that within the first 24 h after release from the female mussel, glochidia viability was

indicative of infectivity, as long as viability was >90%. After 24 h, viability remained relatively

high (>60%) in many species tested, but infectivity of glochidia free in water sometimes declined

rapidly after 24 h. Infectivity was high at all times when viability was >90%. The current

American Society for Testing and Materials (ASTM) guideline for conducting toxicity tests with

2

glochidia of freshwater mussels requires >90% viability for control groups at test termination

(commonly 24 h). Our results indicated that viability is an adequate proxy for infectivity of

glochidia free in water when viability is >90%, and that infectivity decreased markedly in some

species when viability dropped below 90%. Our results also indicated that glochidia released in

conglutinates remained viable and infective for longer periods than glochidia released free into

the water. We also noted for species that brood for several months (long term brooders),

glochidia collected from females toward the end of the brooding season were viable for a shorter

duration than those collected earlier in the brooding season. Based on these data, we recommend

that the current ASTM guideline for conducting toxicity tests with glochidia be retained with a

maximum test duration of 24 h for all mussel species. Additionally, we recommend that

glochidia from long term brooders be collected before the end of the brooding season. For

species with previously defined duration of viability of ≥ 24 h, we recommend a 24 h toxicity

test; however, for species with little or no data on duration of viability, we recommend that

viability be assessed at an intermediate time point(s) (e.g., 6, 12, 18, 24 h) during toxicity tests in

the event that control viability is <90% by 24 h (e.g. at test termination).

In the second phase of the project, we compared the duration of glochidia viability and

infectivity 1) in natural water and sediment versus reconstituted water, and 2) in free glochidia

versus glochidia within conglutinates. These experiments were performed with Ptychobranchus

occidentalis glochidia and rainbow darters (Etheostoma caeruleum). Infectivity of free glochidia

of Ptychobranchus declined to near zero within 24 h, more rapidly than the other species tested.

Viability declined more rapidly in natural water and sediment than in reconstituted water,

perhaps because of microorganisms that attack the glochidia. In contrast to free glochidia,

glochidia in conglutinates remained viable and infective for the duration of the study (96 h). The

3

apparent protective effects of the conglutinate suggest that 24-h toxicity test duration with intact

conglutinates would be conservative.

In a third phase of the project, we evaluated infectivity for glochidia that had been

exposed to NaCl or copper (Cu) for 24 hr. We used glochidia of two mussel species, Lampsilis

cardium and Lampsilis dolabraeformis, and largemouth bass for this study. Two experiments

were performed with glochidia of L. cardium, one with glochidia collected early in the brooding

season (January) and a second with glochidia collected near the end of the brooding season

(June) to determine if maternal brooding time influenced sensitivity to the toxicant, viability, or

infectivity. Following the 24-h exposure to a range of concentrations of NaCl or Cu, we

observed predictable results that viability decreased with increasing toxicant concentration. The

remaining viable glochidia in each treatment were placed on largemouth bass to determine if

they were also able to attach and metamorphose on the fish. In each of the 5 trials, we observed

that the viable glochidia were indeed able to metamorphose at a rate similar to those in the

control. These results suggest that the glochidia viability endpoint is a proxy for metamorphosis

success and is therefore ecologically relevant for 24 h toxicity tests. The data also demonstrated

that glochidia collected early in the brooding season were significantly less sensitive to the

toxicant, had higher metamorphosis success, and remained viable (>90%) longer than those

collected late in the brooding season. These results have important implications for timing of

glochidia collection for toxicity test with long-term brooding mussel species (e.g., members of

the tribe Lampsilini).

In summary, the results of this study provided data that support retaining the current

ASTM guideline for toxicity testing with glochidia using viability as the endpoint, but with

recommended maximum test duration of 24 h. We recommend that glochidia be collected before

4

the end of the brooding season for toxicity tests to ensure health and viability of glochidia. We recommend that glochidia viability be assessed in toxicity tests at a time point between 0 and 24

h (e.g., 6, 12, 18 h) for species for which duration of viability has not been defined in the event

that viability of control mussels of a given species decreases to <90% by the 24 h time point.

Formatted for submission to: Environmental Toxicology and Chemistry

5

Determining the appropriate duration of toxicity tests with glochidia of native

freshwater mussels

Andrea K. Fritts1, M. Christopher Barnhart2, Megan Bradley2, Na Liu2, W. Gregory Cope3, and

Robert B. Bringolf1,4

1 University of Georgia, Warnell School of Forestry & Natural Resources, Athens, Georgia 30602

2 Missouri State University, Department of Biology, Springfield, Missouri 65897

3 North Carolina State University, Department of Environmental and Molecular Toxicology,

Raleigh, North Carolina 27695

4 Corresponding author: [email protected]

6

Abstract

Freshwater mussels (Unionidae) have a remarkable life cycle in which larvae (glochidia) are brooded by the female parent and then released into the water where they must attach to a fish host to metamorphose into a juvenile mussel. Glochidia attach by closing the shell valves on the host. Glochidia also close in response to salt solution, and this closing response to salt is widely used as a measure of viability to determine if glochidia are suitable for host fish suitability trials, as an endpoint for standard toxicity tests, and for a variety of other applications. The lifespan of glochidia in water is limited. We tested the assumption that glochidia viability, measured by the closing response to salt, is indicative of infectivity, the ability of glochidia to attach to a host fish and successfully metamorphose to the juvenile stage. We compared the durations of glochidia viability and infectivity in water for seven mussel species (Lampsilis siliquoidea, Lampsilis cardium, Lampsilis dolabraeformis, Villosa lienosa, Ptychobranchus occidentalis, Amblema plicata, and Utterbackia imbecillis). Viability was indicative of infectivity between 0 and 24 hours. However, often infectivity of glochidia in water decreased after 24 h even when viability remained relatively high, especially for glochidia collected near the end of the brooding season.

The duration of viability and infectivity for glochidia of P. occidentalis was shorter in river water

with sediment compared to reconstituted water. Ptychobranchus occidentalis releases

membrane-bound packets (conglutinates) containing the glochidia. Glochidia in conglutinates

remained infective longer indicating that conglutinates may provide protection from the external

environment for at least four days. We also compared the viability and metamorphosis success of

two mussel species after 24 h exposure to a toxicant (sodium chloride or copper).

Metamorphosis success of remaining viable glochidia did not differ among the concentrations of

7

toxicants, suggesting that viability is indicative of metamorphosis success and that the decrease

in glochidia reproductive potential after exposure to a toxicant is attributed to a smaller number

of glochidia that are open and able to attach to a host, rather than a decrease in metamorphosis.

Our results indicate that glochidia viability is a valid proxy for infectivity and an ecologically

relevant endpoint for standard toxicity tests with freshwater mussels when control viability is

≥90%.

Keywords: Unionidae, larvae, viability, metamorphosis, toxicity

Introduction

Freshwater mussels (Order ) comprise six families and more than 850 species

worldwide (Graf and Cummings 2007). North America is an epicenter of unionid diversity with

approximately 300 species currently recognized (Parmalee and Bogan 1998). The study of

freshwater mussels has increased markedly in recent years, spurred by the recognition that many

mussel taxa have become extinct and many more are threatened by anthropogenic perturbations

(Ricciardi and Rasmussen 1999, Lydeard et al. 2004, Strayer et al. 2004). The extinction or

extirpation of species can have cascading, ecosystem-level effects because freshwater mussels

provide essential ecosystem services such as water filtration, nutrient sequestration and cycling,

and habitat for other aquatic organisms (Vaughn and Spooner 2006, Strayer 2008, Vaughn et al.

2008).

The life cycle of freshwater mussels includes a parasitic stage in which the larvae

(glochidia) must attach to a host (usually a fish) to transform into the juvenile stage. To infect the

8

appropriate host fish species, adult female mussels use a variety of strategies, including mantle

lures, conglutinates, and broadcast of free glochidia into the water (Barnhart et al. 2008, Haag

2012). In each case, the glochidia must be released from the female marsupium and from egg or conglutinate membranes before they are able to contact the host and attach. Attachment occurs

by clamping the valves shut on the host gills or skin. Depending on the host infection strategy,

glochidia may contact the host almost immediately after entering the water or they may remain

free in the water for varying lengths of time before encountering a host.

Lefevre and Curtis (1912) reported that glochidia closed in response to various salt solutions or to blood, and Coker et al. (1921) recommended that the viability of glochidia could be assessed by observing the closing response. This approach remains popular today for assessment of glochidia condition before propagation or host testing. Glochidia that are: a) initially open and b) able to close in response to NaCl are considered to be “viable” and assumed to be able to infect a host fish (Ingersoll et al. 2007). The closing response has also been widely used as an endpoint in toxicological studies (e.g., Keller and Ruessler 1997, Bringolf et al. 2007,

Wang et al. 2007, Cope et al. 2008). In this context, glochidia may either close in response to the toxicant or become unable to close in response to salt. Either condition prevents completion of the lifecycle.

Few quantitative studies have investigated the correlation of the ability of glochidia to close in response to salt (viability) with the ability to attach to a host fish and metamorphose to the juvenile life stage (infectivity). Zimmerman and Neves (2002) assessed effects of different holding temperatures (0°, 10°, 25°C) on glochidia viability and inoculated fish with glochidia

from the 0° and 10°C treatments at 7 and 14 days post extraction from the females.

Metamorphosis to the juvenile stage occurred, but the metamorphosis success (proportion of

9 glochidia that metamorphosed to the juvenile stage) was not reported. Fisher and Dimock (2000) held glochidia of Utterbackia imbecillis in reconstituted water and found that the ability of the glochidia to metamorphose in cell culture media declined more rapidly than the closing response to potassium chloride. To date, it appears that no other studies have quantitatively investigated the relationship between viability and metamorphosis success. With the increasing interest in use of glochidia for standard toxicity tests for derivation of water quality criteria, assessing whether the closing response is indicative of the more ecologically relevant ability of glochidia to successfully attach and metamorphose on a host fish is necessary. A related question is whether the duration of glochidia infectivity in laboratory conditions is similar to that in natural conditions. For example, survival might be shortened by microorganisms present in natural water and sediment but absent from reconstituted water used in laboratory tests. Survival might be lengthened in species whose glochidia are normally protected by conglutinates. Lastly, there is a need to evaluate the effect on metamorphosis of glochidial exposure to a toxicant. Such data would allow researchers to assess if the decrease in reproductive potential is primarily attributed to a smaller number of glochidia that are open and able to attach to a host, or rather a decrease in metamorphosis success.

Our objectives for this study were to determine: 1) if the duration of glochidia viability

(determined by a shell closing response to NaCl) is equivalent to the duration of infectivity--the ability to attach to a host fish and metamorphose successfully into the juvenile stage, 2) the protective effect of conglutinates, 3) the effect of natural water and sediment on glochidia viability and infectivity, and 4) the effect of exposure to a toxicant on metamorphosis success.

These findings will provide insight into the ecological relevance of laboratory tests of glochidia viability, infectivity, and the appropriate duration for standard toxicity tests with glochidia.

10

Methods

Test organisms

We used snorkeling, scuba diving, and tactile searches to collect gravid female mussels from four different states over the course of three years (Table 1). These seven mussel species use the three main infection strategies found in Unionidae: mantle lures (Lampsilis siliquoidea,

Villosa lienosa, Lampsilis cardium, Lampsilis dolabraeformis), conglutinates (Ptychobranchus occidentalis), and glochidia broadcasters (Amblema plicata, Utterbackia imbecillis).

Duration of viability vs. duration of infectivity

We extracted glochidia from 3-8 female mussels per species by gently opening their valves and using a syringe and a gentle stream of water to flush the glochidia from the marsupial gills. Viability of glochidia from individual females was measured by uniformly suspending the glochidia in a known volume of water, removing ten 200-µL subsamples, and placing the subsamples into a 10-cm Petri dish. Each subsample contained ~15-30 glochidia, for a total of approximately 150-300 glochidia tested for viability. A stereomicroscope at 10x magnification was used to count the open and closed glochidia in each of the ten subsamples. A drop of saturated NaCl solution was then added to each 200-µL subsample, and the number of glochidia that remained open 1-min after the addition of the saline solution was counted. Glochidia that were open initially and closed in response to NaCl were considered viable, those that were closed initially were considered functionally dead because they no longer had the ability to attach to a host fish, and those that remained open after the addition of NaCl were considered definitively dead. The viability of a batch of glochidia was calculated by subtracting the number of glochidia

11

that did not close after the addition of NaCl from the number that were originally open, then

dividing by the total number of glochidia (initially open and closed).

Glochidia from female mussels (n= 3-8 with initial viability >90%) were pooled and

randomly divided into replicate (n= 5) glass beakers with 2 L of aerated, moderately hard

reconstituted water (ASTM 2006a) where they were maintained at 20°C (± 1oC) with a 16L: 8D

photoperiod. At the beginning of the experiment and every 24 h thereafter, the viability of each

replicate was calculated following the method described above. Subsampling continued daily

until mean viability for the five replicates was <10%. At each time point, a second subsample of glochidia from each replicate was tested for infectivity on known host fish (two fish per

replicate) for a total of 10 fish per time point. Host fish, largemouth bass (Micropterus

salmoides) (8-18 cm) and rainbow darters (Etheostoma caeruleum) (5-8 cm) were obtained from hatcheries, which insured that the fish had not been previously exposed to glochidial infections.

Host fish were exposed to 4000 viable glochidia per liter of water for 15 min; glochidia were kept in suspension with a large-bulb pipette and vigorous aeration.

Attachment and metamorphosis success were assessed in multiunit recirculating aquarium systems (Aquatic Habitats, Apopka, Florida) as described by Dodd et al. (2005). Fish infested with glochidia were rinsed, then placed individually in unit tanks (1-3 L) fitted with 153­

μm filters to collect all sloughed glochidia and metamorphosed juveniles. Juveniles were distinguished from glochidia by possessing a foot and valve movement. Contents of the filters were examined under a stereomicroscope every 1-2 days for a minimum of 14 days and until no glochidia or juveniles were collected for at least three consecutive days. Percent metamorphosis was calculated by dividing the total number of juveniles by the total number of glochidia and juveniles recovered from each fish. The aquarium systems were maintained at 20-22°C. All

12 water-chemistry variables were measured daily throughout the study and ranged from 7.5 to8.2 mg/L dissolved oxygen, pH: 6.8 to7.6, and total ammonia nitrogen <0.1 mg/L (Table 2).

Effects of conglutinates and sediment

The durations of glochidia viability and infectivity in conglutinates and free in water were compared in Ptychobranchus occidentalis. These durations in reconstituted water and in natural water and sediment were also compared. Hosts were rainbow darter (Etheostoma caeruleum, 5-8 cm total length). Conglutinates were dissected from the marsupial demibranchs of gravid female mussels with No. 0 insect pins. Conglutinates from six females were pooled, and each conglutinate was examined carefully under magnification to determine if it was intact.

Only unbroken conglutinates were used for conglutinate exposures. Free glochidia were obtained by using pins to gently tear open the outer conglutinate membrane. The ruptured conglutinate was then gently drawn in and out of a 3-mL transfer pipette to dislodge the glochidia.

Both free glochidia and intact conglutinates were exposed to reconstituted hard water or to river water with fine sediment. Reconstituted water was similar in formula to that of Smith et al. (1997), but concentrations of all solutes were doubled. River water and wet sediment were collected from the James River (Greene County, Missouri). Approximately 15L of water and

250-ml of sediment were combined and filtered to 32 µm with Nitex filter cloth. Water and sediment were homogenized by stirring with a paint stirrer while being portioned in 200-ml aliquots to each of the test chambers (250-ml beakers). A layer of sediment about 1 mm deep settled in each beaker. Treatments (three replicates per treatment x exposure time; 60 beakers total) were: 1) reconstituted water with free glochidia, 2) reconstituted water with conglutinates,

3) river water and sediment with free glochidia, and 4) river water and sediment with

13 conglutinates. Each beaker received 1000 glochidia or 3 conglutinates containing a similar total number of glochidia. At time zero, and then each 24-h interval for 96 h, viability and infectivity were assessed. In conglutinate treatments, the conglutinates were opened and the glochidia freed. Two darters were placed in each beaker for 15 min, while gently stirring the water with a pipette, to achieve infestation. Methods for calculating viability and infectivity were similar to those previously described.

Effects of glochidial exposure to toxicants

We extracted glochidia from 3-8 female mussels per species by gently opening their valves and using a syringe and a gentle stream of water to flush the glochidia from the marsupial gills. Viability of glochidia from individual females was measured in the same manner as that described above. According to ASTM guidelines (ASTM 2006b), glochidia from female mussels (n= 3-8 with initial viability >90%) were pooled and randomly divided into six concentrations of NaCl (0, 0.5, 1.0, 1.5, 2.0, 2.5 ppt) or four concentrations of copper (0, 15, 30,

60 ppm) dissolved in moderately hard reconstituted water (ASTM 2006a). In all glochidia exposures temperature was 20oC ±1oC, pH ranged from 7.1 to 7.5, dissolved oxygen was >80%

of saturation, hardness was 75-83 mg/L as CaCO3, and alkalinity was 71-85 mg/L as CaCO3

(Table 3, Table 4). Glochidia from each concentration were then split into replicate (n= 3) glass

beakers where they were maintained at 20°C with a 16L: 8D photoperiod. After 24 h of exposure

to the respective toxicants, the viability of each replicate was calculated following the method

described above. A second subsample of glochidia from each replicate was tested for infectivity

on largemouth bass (two fish per replicate) for a total of six fish per concentration. The

inoculation procedure follows the methods described above. In each replicate, the inoculation

14 suspension was adjusted to provide 4000 viable glochidia per liter. The aquarium systems were maintained at 18-22°C. All water-chemistry variables were measured daily throughout the study and ranged from 7.5 to 8.6 mg/L dissolved oxygen, pH: 6.8 to7.7, and total ammonia nitrogen

<0.1 mg/L. Concentrations of NaCl were verified daily in one replicate of each treatment with a handheld salinity meter (YSI 30, Yellow Springs Instruments, Yellow Springs, OH) and were within 0.1 ppt of nominal at all times. Water samples were collected and preserved to pH <2 for

Cu analysis prior to analysis by inductively coupled plasma mass spectrometry. All measured Cu concentrations (n=10) were 80-120% of nominal and duplicate samples (n=4) varied by <5%.

All statistical analyses were based on nominal concentrations.

Statistical analysis

Data were tested for normality using the Kolmogorov-Smirnov test (SAS, version 9.3;

SAS Institute, Cary, North Carolina). To achieve normality and equal variance, viability and metamorphosis data were transformed with the arcsine of the square root. Transformed data were analyzed with a one-way ANOVA, followed by a post-hoc Dunnett’s test to assess for differences between the control (i.e., 0 h in the aging exposures) and each time point or toxicant concentration. Attachment data (the number of glochidia attached per fish) were analyzed with a one-way ANOVA, followed by a post-hoc Tukey’s test. Level of significance for all tests was set at α = 0.05. A 24-h EC50 was calculated for each 24-h glochidia viability test with the

Trimmed Spearman-Karber Method (ToxStat, WEST, Inc.). We considered EC50s significantly different based on non-overlapping 95% confidence intervals (Bringolf et al. 2007).

15

Results

Duration of viability vs. duration of infectivity

The initial (0 h) glochidia viability met the current American Society for Testing and

Materials (ASTM) guideline of >90% (ASTM 2006b) for all seven mussel species tested;

however, the trial with L. siliquoidea collected in June had an initial viability of only 80%.

Viability and metamorphosis remained higher for longer with L. siliquoidea glochidia collected

in January (Fig. 1A) compared to those collected in June (Fig. 1B), which is the end of the

brooding season for many members of the tribe Lampsilini. At the 24-h time point, viability was

<90% for L. cardium, U. imbecillis, and L. siliquoidea (June) (Fig. 1). According to the ASTM

guideline for conducting laboratory toxicity tests with freshwater mussels (ASTM 2006b), data

from these time points would not be valid for toxicological testing purposes because the viability

was <90%. Five species (L. siliquoidea (Jan), L. dolabraeformis, V. lienosa, A. plicata, and P. occidentalis) had >90% viability at 24 h (Fig. 1). For these species, metamorphosis success did not differ between 0 and 24 h (Fig. 1, Table 4). The exception was P. occidentalis free glochidia, which had viability >90% through 72 h but had low (<5%) metamorphosis by 24 h

(Fig. 2A). For all but one species, metamorphosis success decreased rapidly as viability dropped below 90%, although viability remained near or above 60% for 3 d with four of the seven species. Overall, metamorphosis success ranged from 19 to 78% (mean 54%) at time zero for the species tested, typical of rates achieved in mussel propagation trials with known host fish-mussel pairs, at common temperatures, and with naïve test fish (Dodd et al. 2005). Total glochidia attachment did not differ among treatments for any of the mantle lure or conglutinate species tested (p = 0.08-0.64), but the two broadcasting species (U. imbecillis and A. plicata) had significantly greater glochidia attachment (p = <0.001) at the final inoculation time point (mean

16 attachment ± 95% CI) (U. imbecillis: 0 to24 h = 353 ± 61, 48 h = 1306 ± 208; A. plicata: 0 to 48 h = 365 ± 44, 72 h = 745 ± 88).

Effects of conglutinates and sediment

Initial (0 h) glochidia viability was > 90% in all treatments. Viability of free glochidia exposed to sediment and river water decreased rapidly (52% viable by 48 h) whereas viability of free glochidia in reconstituted water (no sediment) remained >90% until 72 h (Fig. 2A & 2C).

Sediment also reduced the duration of viability for glochidia in conglutinates but viability did not decrease to <90% until 72 h of exposure (Fig. 2B & 2D). Viability of conglutinates in reconstituted water was >90% for at least 96 h, the time at which the experiment was terminated.

Metamorphosis success of free glochidia declined more rapidly than that of glochidia within conglutinates and declined similarly in both river water and reconstituted water (Fig. 2,

Table 2). Metamorphosis success was high (45% ± 13) for glochidia placed on fish immediately after removal from the mussel, but decreased rapidly to <5% for free glochidia aged 24 h or more, even though viability remained high (>90%). In contrast, glochidia in conglutinates remained infective for at least 96 h. Total glochidia attachment (mean ± 95% CI) did not differ among P. occidentalis conglutinate treatments nor free glochidia in reconstituted water (38 ± 6; p

= 0.11-0.61), but free glochidia exposed to river water and sediment had significantly less glochidia attachment (p = 0.0113) during all inoculation time points after the initial inoculation

(0 h = 45 ± 20; 24-96 h = 12 ± 3).

Effects of glochidial exposure to toxicants

17

In each toxicant test, viability decreased (ranged from 95% to 25%) as toxicant

concentration increased (Fig. 3). However, for the remaining viable glochidia in each treatment,

metamorphosis success was similar among treatments regardless of viability (Fig. 3, Table 5).

Lampsilis cardium glochidia collected in December (early brooding period; Fig. 3A) maintained

high viability longer than those of the same species collected in June (late brooding period; Fig.

3B) and also had consistently higher (55-70%) metamorphosis rates than those collected in June

(18-23%). Glochidia collected early in the brooding period (December) were significantly less

sensitive (based on non-overlapping 95% confidence intervals) to NaCl than those collected late

in the brooding period (June). The 24-h EC50 (± 95% CI) for December glochidia was 2.04 ±

0.11 and for June glochidia was 0.80 ± 0.16 (Fig. 3D). Total glochidia attachment (mean ± 95%

CI) did not differ among L. cardium or L. dolabraeformis trials (p = 0.16-0.58).

Discussion

Our results indicate that glochidia viability (as measured by the shell closing response to

NaCl exposure) was indicative of the ability to metamorphose into the juvenile life stage, as long as viability was >90%. Timing of glochidia collection also appears to be critical to glochidia health; viability of glochidia collected late in the brooding season declined rapidly compared to those collected earlier in the brooding season. The current ASTM guideline for conducting toxicity tests with glochidia of freshwater mussels (ASTM 2006b) requires >90% viability for control groups at test termination (commonly 24 h) and our data support the continued use of this criteria. Additionally, glochidia that remained viable following exposure to a toxicant for 24 h were able to successfully metamorphose, thus the viability endpoint appears to have direct ecological relevance. Based on these data, we recommend that the current ASTM guideline for

18

glochidia toxicity testing be retained with a minimum control viability of 90% throughout the

duration of the test for all mussel species. Additionally, we recommend that glochidia from long

term brooders be collected before the end of the brooding season because those collected late in

the season appeared less healthy based on our results that showed glochidia collected late in the

brooding season were more sensitive to toxicants, had lower initial viability which declined

quickly, and had poorer metamorphosis success. For species with previously defined duration of

viability of ≥ 24 h, we recommend a 24 h toxicity test; however, for species with little or no data

on duration of viability, we recommend that viability be assessed at an intermediate time point(s)

(e.g., 6, 12, 18, 24 h) during toxicity tests in the event that control viability is <90% by 24 h (e.g.

at test termination). Shipping time must also be a consideration if glochidia cannot be obtained

locally. Further, for species with previously defined viability of ≥90% for ≥24h, tests of shorter duration (because of low control survival) should not be considered acceptable.

The duration of viability for free glochidia of P. occidentalis was shorter in river water and sediment than in reconstituted water, but the effect was much less pronounced for glochidia in conglutinates. We suspect that the relatively rapid decline of free glochidia in river water with sediment was the result of microorganisms that compromised glochidia survival and ability to attach to a host fish. In a preliminary test of this hypothesis, a group of free glochidia was tested in 0.4 µ filter-sterilized river water and they remained >80% viable at 72 h. This result indicated that microorganism removal positively affected viability. The subsequent infectivity of these glochidia was not tested, however. Whereas infectivity of free glochidia of P. occidentalis

decreased rapidly in both treatments, infectivity of glochidia in conglutinates remained high for

the duration of the study (96 h). The apparent protective effects of the conglutinate suggest that a

24-h toxicity test duration would be conservative for species that release glochidia in

19

conglutinates. Conglutinates provide protection from some toxicants (e.g., copper) and thereby

reduce toxicity (increase LC50, EC50 values; Gillis et al. 2008).

Our results suggest that the conglutinate packet of P. occidentalis provides protection

from the external environment for at least 4 d. Even greater protection may be afforded by the

marsupial gill of the brooding female, where glochidia of long-term brooders develop and

survive for many months (Haag 2012). Once released from the protection of the conglutinate or

the marsupia of the female mussel, the infectivity of free glochidia appears to decrease after 24 h, depending on the species. Our data indicate that metamorphosis success decreases before viability decreases; at 48 h post extraction, L. siliquoidea (Jan) and V. lienosa exhibited 80% viability, but the corresponding metamorphosis success was reduced by 50% (Fig. 1A &1B).

Importantly, the reduced infectivity was not a result of lower attachment to host fish, which remained consistent across inoculation days.

The copper and NaCl toxicant exposure results suggest that the decrease in glochidia reproductive potential after exposure to a toxicant is attributed to a smaller number of glochidia that are open and able to attach to a host, rather than a decrease in metamorphosis success. Since the inoculation suspensions were adjusted to a uniform number of viable glochidia, the consistent metamorphosis success supports the use of the viability test as a proxy for metamorphosis.

These results were consistent with 2 mussel species and two toxicants with different modes of action.

For mussel species that are long-term brooders, the length of time that glochidia have aged within the female mussel also may affect glochidia function and condition. Our data suggest that older glochidia may be in poorer physiological condition (e.g., less energy reserves) and are unable to remain viable for as long after the glochidia are removed from the female. In

20 our study, L. siliquoidea glochidia early in the brooding period (January) registered 80% viability 48-h after being removed from the female, whereas glochidia from the same species collected late in the brooding period (June) only had 20% viability at 48 h post extraction.

Additionally, Lampsilis cardium glochidia tested late in the brooding period (June) also showed a rapid decline in viability compared to L. cardium tested early in the brooding period

(December). Dodd et al. 2006 reported that Lampsilis reeveiana glochidia aged 241-360 d had significantly less metamorphosis success (67.5%) compared to glochidia aged 0-120 d (87.5%) and 121-240 d, (88.7%). Cope et al. (2008) reported longer duration of viability (>90%) for some of the species used in the present study; however, timing of collection of the glochidia was not reported.

The implications for the results of this study are multifaceted. For the purposes of toxicological testing, our recommendation is to retain the current ASTM guideline for toxicity testing with glochidia, but specify a maximum test duration of 24 h when using viability (i.e., the valve closure response to NaCl) as the endpoint. Ideally toxicity tests should be conducted with glochidia immediately following removal from the female mussel, thus shipping of brooding females rather than free glochidia may be considered. However, shipping of adult mussels is less practical and may create logistical and regulatory (e.g., interbasin transfer of species or threatened or endangered status) problems, so shipping of free glochidia may be necessary. Our results indicate that this time frame may be suitable (i.e., viability and infectivity can remain

≥90% for 48 h) but we recommend that viability be assessed at one or more time points between

0 and 24 h (e.g., 6, 12, 18 h) in the event that viability of controls decreases to <90% by the 24 h time point of the toxicity test. This approach is especially important when a species is being used for which no viability duration information has been previously reported. These

21 recommendations are supported by Cope et al. (2008), who showed that viability (determined with the NaCl method) of glochidia from 20 species of mussels, three of which were tested in this study (L. cardium, L. siliquoidea, U. imbecillis), was suitable (≥90%) for 24 h or less. This time period corresponds with the ecologically relevant endpoint of infectivity determined in this study. The higher toxicant sensitivity, lower initial viability and lower metamorphosis success of glochidia collected late in the brooding season suggests they are not appropriate for use in toxicity tests. Use of glochidia from late in the brooding period (for long-term brooders) may result in toxicity test results that are substantially different that if glochidia were collected early in the brooding season. More research is required to better define the period when glochidia fitness begins to decline to the point that toxicant sensitivity is increased. Furthermore, implications for freshwater mussel host suitability research include the possibility of obtaining a false-negative result, if the glochidia have been outside of the female for more than 24 h. The decrease in metamorphosis success should be noted by those who conduct propagation efforts with imperiled mussels, and every effort should be made to place glochidia onto host fish as soon as practical after the glochidia are removed from the female mussel. Future propagation and culture research evaluating the infectivity of the glochidia life stage of freshwater mussels would benefit from accurate calculation and reporting of viability data.

In summary, we found that glochidia viability of ≥90% was indicative of infectivity; however, after 24 h infectivity often declined more rapidly than viability. Glochidia of long-term brooders collected late in the brooding season had low fitness compared to those collected early in the brooding season. Glochidia in contact with sediment may have a reduced duration of viability and infectivity, but glochidia in conglutinates may remain viable and infective for extended periods after release from brooding females. Glochidia viability is, therefore, an

22

ecologically relevant endpoint in standard toxicity tests when viability is high and the current

ASTM criteria for control viability (90%) is appropriate.

Acknowledgements

Funding for this research was provided by the U.S. Environmental Protection Agency Region 5

Great Lakes Restoration Initiative Program through Work Order no. 104 from the U.S.

Geological Survey. We thank Charles Delos, Ed Hammer, Linda Holst, Lisa Huff, Dave Mount,

Charles Stephan, and Brian Thompson of the U.S. EPA for their valuable insight and assistance

on project need and experimental design. We thank J. Creamer, M. Fritts, R. Ratajczak, A.

Tuck, and J. Wisniewski for assistance in the laboratory and field, and B. Sietman and T.

Newton for providing mussels. Largemouth bass were donated by Owens and Williams Fish

Hatchery (Hawkinsville, Georgia) and rainbow darters were provided by Conservation Fisheries

(Knoxville, Tennessee). The Georgia Cooperative Fish and Wildlife Research Unit is jointly

supported by the University of Georgia, Georgia Department of Natural Resources, U.S.

Geological Survey, U.S. Fish and Wildlife Service, and Wildlife Management Institute. Any use

of trade, product, or firm names is for descriptive purposes only and does not imply endorsement

by the U.S. Government.

23

Literature cited

ASTM (American Society for Testing and Materials). 2006a. Standard guide for conducting

acute toxicity tests on test materials with fishes, macroinvertebrates, and amphibians.

E729-96. pp 79–100 in Annual Book of ASTM Standards, Vol 11.06. Philadelphia, PA.

ASTM (American Society for Testing and Materials). 2006b. Standard guide for conducting

laboratory toxicity tests with freshwater mussels E2455-06. pp 1393–1444 in Annual

Book of ASTM Standards, Vol 11.06. Philadelphia, PA.

Barnhart, M. C., W. R. Haag, and W. N. Roston. 2008. Adaptations to host infection and larval

parasitism in Unionoida. Journal of the North American Benthological Society 27:370­

394.

Bringolf, R. B., W. G. Cope, M. C. Barnhart, S. Mosher, P. R. Lazaro, and D. Shea. 2007.

Acute and chronic toxicity of pesticide formulations (atrazine, chlorpyrifos and

permethrin) to glochidia and juveniles of Lampsilis siliquoidea (Unionidae).

Environmental Toxicology and Chemistry 26:2101–2107.

Coker, R. E., A. F. Shira, H. W. Clark, and A. D. Howard. 1921. Natural history and propagation

of freshwater mussels. Bulletin of the U.S. Bureau of Fisheries 37:75–182.

Cope, W. G., R. B. Bringolf, D. B. Buchwalter, T. J. Newton, C. G. Ingersoll, N. Wang, T.

Augspurger, F. J. Dwyer, M. C. Barnhart, R. J. Neves, and E. Hammer. 2008. Differential

exposure, duration, and sensitivity of unionoidean bivalve life stages to environmental

contaminants. Journal of the North American Benthological Society 27:451-462.

Dodd, B. J., Barnhart, M.C., Rogers-Lowery, C.L., Fobian, T.B. & R.V. Dimock Jr. 2005. Cross

resistance of largemouth bass to glochidia of unionid mussels. Journal of Parasitology

91:1064-1072.

24

Dodd, B. J., M. C. Barnhart, C. L. Rogers-Lowery, T. B. Fabian, and R. V. Dimock. 2006.

Persistence of host response to glochidia larvae in Micropterous salmoides. Fish and

Shellfish Immunology 21:473–484.

Fisher, G. R. and R. V. Dimock. 2000. Viability of glochidia of Utterbackia imbeccilus

(: Unionidae) following their removal from the parent mussel. Pages 185-188 in

R. A. Tankersley, D. I. Warmolts, G. T. Watters, B. J. Armitage, P. D. Johnson, and R. S.

Butler (editors). Freshwater Mollusk Symposia Proceedings. Ohio Biological Survey,

Columbus, Ohio. 241 pp.

Graf, D. L. and K. S. Cummings. 2007. Review of the systematics and global diversity of

freshwater mussel species (Bivalvia : Unionoida). Journal of Molluscan Studies 73:291­

314.

Gillis, P. L., R. J. Mitchell, A. N. Schwalb, K. A. McNichols, G. L. Mackie, C. M. Wood, and J.

D. Ackerman. 2008. Sensitivity of the glochidia (larvae) of freshwater mussels to copper:

assessing the effect of water hardness and dissolved organic carbon on the sensitivity of

endangered species. Aquatic Toxicology 88(2):137-145.

Haag, W. R. 2012. North American Freshwater Mussels: Natural History, Ecology, and

Conservation. Cambridge University Press, Cambridge, United Kingdom. 536 pp.

Ingersoll, C. G., N. J. Kernaghan, T. S. Gross, C. D. Bishop, N. Wang, and A. Roberts. 2007.

Laboratory toxicity testing with freshwater mussels. Pages 95-134 in J. L. Farris and J. H.

Van Hassel (editors). Freshwater bivalve ecotoxicology. CRC Press, Boca Raton, Florida,

and SETAC Press, Pensacola, Florida. 321 pp.

25

Keller, A. E. and D. S. Ruesller. 1997. The toxicity of malathion to unionid mussels: relationship

to expected environmental concentrations. Environmental Toxicology and Chemisty

16(5):1028-1033.

Lefevre, G. and W. C. Curtis. 1912. Studies on the reproductive and artificial propagation of

fresh-water mussels. Bulletin of the Bureau of Fisheries 30:105-201+12 plates.

Lydeard, C., R. H. Cowie, W. F. Ponder, A. E. Bogan, P. Bouchet, S. A. Clark, K. S. Cummings,

T. J. Frest, O. Gargominy, D. G. Herbert, R. Hershler, K. E. Perez, B. Roth, M. Seddon,

E. E. Strong, and F. G. Thompson. 2004. The global decline of nonmarine mollusks.

Bioscience 54:321-330.

Parmalee, P. W. and A. E. Bogan. 1998. The Freshwater Mussels of Tennessee. University of

Tennessee Press, Knoxville, TN. 328 pp.

Ricciardi, A. and J. B. Rasmussen. 1999. Extinction rates of North American freshwater fauna.

Conservation Biology 13:1220-1222.

Smith, M. E., J. M. Lazorchak, L. E. Herrin, S. Brewer-Swartz, and W. T. Thoeny. 1997. A

reformulated, reconstituted water for testing the freshwater amphipod, Hyalella azteca.

Environmental Toxicology and Chemistry 16(6):1229-1233.

Strayer, D. L., J. A. Downing, W. R. Haag, T. L. King, J. B. Layzer, T. J. Newton, and S. J.

Nichols. 2004. Changing perspectives on pearly mussels, North America's most imperiled

. Bioscience 54:429-439.

Strayer, D. L. 2008. Freshwater Mussel Ecology: A Multifactor Approach to Distribution and

Abundance. University of California Press, Berkeley, California. 204 pp.

26

Vaughn, C. C., and D. E. Spooner. 2006. Unionid mussels influence macroinvertebrate

assemblage structure in streams. Journal of the North American Benthological Society

25:691–700.

Vaughn, C. C., S. J. Nichols, and D. E. Spooner. 2008. Community and foodweb ecology of

freshwater mussels. Journal of North American Benthological Society 27:409–423.

Wang, N., C. G. Ingersoll, D. K. Hardesty, C. D. Ivey, J. L. Kunz, T. W. May, F. J. Dwyer, A. D.

Roberts, T. Augspurger, C. M. Kane, R. J. Neves, and M. C. Barnhart. 2007. Acute

toxicity of copper, ammonia, and chlorine to glochidia and juveniles of freshwater

mussels (Unionidae). Environmental Toxicology and Chemistry 26:2036–2047.

Zimmerman, L. L. and R. J. Neves. 2002. Effects of temperature on duration of viability for

glochidia of freshwater mussels (Bivalvia: Unionidae). American Malacological Bulletin

17:31-35.

27

Table1: Dates and locations of freshwater mussel collections.

Species Water body County Date Amblema plicata Sac River Cedar, MO Jul 2011 Villosa lienosa Cooleewahee Creek Baker, GA Oct 2010 Lampsilis cardium Mississippi River La Crosse, WI Jun 2011 L. siliquoidea Silver Fork of Perche Creek Boone, MO Jan 2011 L. siliquoidea Red Lake River Pennington, MN Jun 2012 L. dolabraeformis Oconee River Montgomery, GA Oct 2012 Utterbackia imbecillis Lake Oconee Morgan, GA Jul 2011 Ptychobranchus occidentalis St. Francis River Wayne, MO Feb 2011

Table 2. Water quality parameters (mean ± SD) for ASTM moderately hard reconstituted water used for glochidia viability and infectivity studies in which the glochidia were aged before inoculation on fish. Hardness and alkalinity (mg CaCO3/L) were measured once at the beginning of each study, other parameters were measured at least 10 times throughout each study.

Dissolved Temp Hardness Alkalinity Oxygen (oC) (mg CaCO3/L) (mg CaCO3/L) Study pH (mg/L) Amblema plicata 7.1 ± 0.2 7.7 ± 0.9 20.5 ± 0.2 75 55 Villosa lienosa 7.7 ± 0.2 8.1 ± 1.0 19.7 ± 0.1 85 62 Lampsilis cardium 7.5 ± 0.2 7.6 ± 0.9 20.4 ± 0.1 82 64 L. siliquoidea (Jan) 7.6 ± 0.1 7.8 ± 1.0 19.4 ± 0.2 80 62 L. siliquoidea (June) 7.6 ± 0.2 8.1 ± 0.9 20.3 ± 0.1 82 60 L. dolabraeformis 7.7 ± 0.1 7.9 ± 0.8 19.9 ± 0.1 84 65 Utterbackia imbecillis 7.2 ± 0.1 7.1 ± 1.0 19.4 ± 0.1 78 56

28

Table 3. Water quality parameters for ASTM moderately hard reconstituted water used for glochidia viability and infectivity studies in which the glochidia were exposed to a gradient of

NaCl for 24 hours, after which time fish were inoculated. Hardness and alkalinity (mg CaCO3/L) was measured at the beginning of the exposure and all other parameters were measured at the end of the 24 hour exposure.

NaCl (ppt) Study 0 0.5 1.0 1.5 2.0 Lampsilis cardium (Dec) pH 7.3 7.9 7.9 7.9 7.9 DO (mg/L) 6.3 6.0 6.3 6.2 6.4 Temp (°C) 18.4 18.6 18.5 18.6 18.7 Hardness 86 86 86 86 86 Alkalinity 62 62 62 62 62 L. cardium (June) pH 7.0 7.3 7.3 7.3 7.3 DO (mg/L) 8.1 8.3 5.7 5.4 6.6 Temp (°C) 19.4 19.3 19.4 19.4 19.7 Hardness 88 88 88 88 88 Alkalinity 64 64 64 64 64 L. dolabraeformis pH 7.6 7.7 7.6 7.7 7.8 DO (mg/L) 8.4 8.2 7.8 8.1 7.4 Temp (°C) 20.9 20.8 20.8 20.9 20.8 Hardness 80 80 80 80 80 Alkalinity 60 60 60 60 60

Table 4. Water quality parameters for ASTM moderately hard reconstituted water during Lampsilis dolabraeformis glochidia viability and infectivity test in which the glochidia were exposed to a gradient of copper (Cu) for 24 hours, after which time fish were inoculated. Hardness and alkalinity (mg CaCO3/L) was measured at the beginning of the exposure and all other parameters were measured at the end of the 24 hour exposure.

Cu (ppm) Parameter 0 15 30 60 pH 7.7 7.7 7.5 7.6 DO (mg/L) 8.7 7.1 8.2 7.9 Temp (°C) 20.9 20.9 20.8 20.7 Hardness 80 80 80 80 Alkalinity 60 60 60 60

29

Table 5. Results of one-way ANOVA comparing the change in glochidia viability or metamorphosis over time for seven freshwater mussel species. Degrees of freedom (df), test statistic (F value) and P values are presented. Level of significance was α = 0.05.

Species df F value P value Viability Amblema plicata 4, 20 45.23 <0.0001 Villosa lienosa 5, 24 59.83 <0.0001 Lampsilis cardium 3, 16 211.85 <0.0001 L. siliquoidea, (Jan) 5, 24 47.13 <0.0001 L. siliquoidea, (June) 2, 12 31.69 0.0005 L. dolabraeformis 4, 20 75.39 <0.0001 Utterbackia imbecillis 3, 16 286.73 <0.0001 Ptychobranchus occidentalis, glochidia/water 4, 10 13.18 0.0005 P. occidentalis, conglutinate/water 4, 10 0.83 0.5372 P. occidentalis, glochidia/sediment 4, 10 35.17 <0.0001 P. occidentalis, conglutinate/sediment 4, 10 4.00 0.0344 Metamorphosis A. plicata 3, 16 7.75 0.0020 V. lienosa 3, 16 9.25 0.0009 L. cardium 2, 12 49.65 <0.0001 L. siliquoidea, (Jan) 4, 20 86.85 <0.0001 L. siliquoidea, (June) 1, 8 2.68 0.1404 L. dolabraeformis 3, 16 20.41 <0.0001 U. imbecillis 2, 12 105.09 <0.0001 P. occidentalis, glochidia/water 4, 10 32.69 <0.0001 P. occidentalis, conglutinate/water 4, 10 6.68 0.0070 P. occidentalis, glochidia/sediment 4, 10 6.77 0.0066 P. occidentalis, conglutinate/sediment 4, 10 1.37 0.3118

30

Table 6. Results of one-way ANOVA comparing the change in glochidia viability or metamorphosis for glochidia exposed to increasing concentrations of sodium chloride (NaCl) or copper (Cu). Degrees of freedom (df), test statistic (F value) and P values are presented. Level of significance was α = 0.05.

Species df F value P value Viability L.cardium (Dec), NaCl 5, 12 45.65 <0.0001 L.cardium (Jun), NaCl 3, 8 165.67 <0.0001 U.imbecillis, NaCl 2, 6 58.49 0.0001 L.dolabraeformis, NaCl 5, 12 92.31 <0.0001 L.dolabraeformis, Cu 3, 8 188.06 <0.0001 Metamorphosis L.cardium (Dec), NaCl 5, 12 0.82 0.5595 L.cardium (Jun), NaCl 3, 8 2.67 0.1187 U.imbecillis, NaCl 2, 6 3.73 0.0886 L.dolabraeformis, NaCl 5, 12 10.21 0.0005 L.dolabraeformis, Cu 3, 8 3.13 0.0875

31

Figure Legends

Fig. 1. Comparison of percent viability (bars) and percent metamorphosis success (lines; ± 95% CI) for glochidia following release from female mussels of A) Lampsilis siliquoidea (Jan), B) Lampsilis siliquoidea (Jun), C) Lampsilis cardium, D) Lampsilis dolabraeformis, E) Amblema plicata, F) Villosa lienosa, and G) Utterbackia imbecillis. Viability was determined with approximately 200 glochidia per replicate (n=5) and metamorphosis success was assessed on largemouth bass (n=10 fish, two per replicate). Presence of a black asterisk (*) indicates a significant difference in viability, and a white asterisk indicates a significant difference in metamorphosis success.

Fig. 2. Comparison of percent viability (bars) and percent metamorphosis success (lines; ± 95% CI) for glochidia of Ptychobranchus occidentalis: A) free glochidia in reconstituted water, B) glochidia in conglutinates in reconstituted water, C) free glochidia exposed to river water and sediment, and D) glochidia in conglutinates exposed to river water and sediment. Viability was determined with approximately 200 glochidia per replicate (n=3) and metamorphosis success was assessed on rainbow darter (n=6 fish, two per replicate). Presence of a black asterisk (*) indicates a significant difference in viability, and a white asterisk indicates a significant difference in metamorphosis success.

Fig. 3. Comparison of percent viability (bars) and percent metamorphosis success (lines; ± 95% CI) for glochidia exposed to increasing concentrations of sodium chloride (NaCl) or copper (Cu) for 24 h: A) Lampsilis cardium (Dec) NaCl exposure, B) Lampsilis cardium (Jun) NaCl exposure, C) Lampsilis dolabraeformis exposed to Cu. Viability was determined with approximately 200 glochidia per replicate (n=3) and metamorphosis success was assessed on largemouth bass (n=6 fish, two per replicate). Presence of a black asterisk (*) indicates a significant difference in viability, and a white asterisk indicates a significant difference in metamorphosis success.

Fig. 4. Comparison of NaCl 24-h EC50 (± 95% confidence interval) for Lampsilis cardium glochidia collected early in the brooding period (December) and late in the brooding period (June). Asterisk indicates significant difference in EC50s (non-overlapping 95% confidence intervals).

32

Fig. 1. A 100 * 80 * 60 * 40 * * * Percent (%) 20 * 0 0 2448 72 96 120 Exposure (h) B 100 80 * 60 40 * Percent (%) 20 0 0 2448 Exposure (h)

C 100 * 80 60 * 40 20 * Percent (%) * * 0 0 2448 72 Exposure (h)

33

D 100 * 80 * 60 40

Percent (%) 20 * * 0 0 2448 72 96 Exposure (h) E 100 * 80 * 60 40 *

Percent (%) 20 * * * 0 0 2448 72 96 120 Exposure (h)

F 100 80 * 60 * 40

Percent (%) 20 * * * 0 0 24 48 72 96

Exposure (h)

34

G 100 80 60 * 40 *

Percent (%) * 20 * 0 * 024 48 72 Exposure (h)

35

Fig. 2. A 100 * 80 * 60 40

Percent (%) 20 * * * * 0 0 24 48 72 96

Exposure (h) B 100 80 * 60 * 40

Percent (%) 20 0 0 2448 72 96 Exposure (h) C 100 80 * 60 * 40 * Percent (%) 20 * * 0 * * 0 2448 72 96 Exposure (h)

36

D 100 80 60 * 40

Percent (%) 20 0 0 2448 72 96

Exposure (h)

37

Fig. 3. A 100 * * 80 * 60 40

Percent (%) 20 0 0.0 0.5 1.0 1.5 2.0 NaCl (ppt) B 100 80 * 60 * 40 *

Percent (%) 20 * 0 0.0 0.5 1.0 1.5 2.0 NaCl (ppt) C 100 * 80 * 60 * * 40 *

Percent (%) 20 0 015 30 60

Copper (ppm)

38

Fig. 4. 2.5 2.0 1.5 1.0 * 0.5 24-h EC50 (g/L) 24-h EC50 0.0 Dec June

39