Estrogens, Endocrine Disruption, and Approaches to Assessing Gametogenesis and Reproductive Condition in Freshwater Mussels (Bivalvia: Unionidae)

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy In the Graduate School of The Ohio State University

David M. Sovic, B.S. Graduate Program in Environmental Science The Ohio State University 2016

Dissertation Committee: Roman P. Lanno, Advisor G. Thomas Watters Susan Fisher Linda Weavers

Abstract

Organisms belonging to the family Unionidae, commonly referred to as freshwater mussels, unionids, or pearly mussels, have, over the last several decades, experienced drastic declines in range and number. These declines have not been localized to a particular region and have not been specific to any particular environment or habitat, but have been experienced by a great number of the species belonging to this diverse group. While a variety of potentially causative factors have been implicated in the decline of Unionids worldwide, the possibility that environmental contaminants, including those identified or suspected as endocrine disrupting chemicals (EDCs), might contribute to observed unionid declines is central to this work. Unionid bivalves are long-lived, sessile creatures that, by actively filter feeding, are at risk of a high degree of exposure to water-borne xenobiotics. Ecotoxicological research on potential endocrine disruptive effects on bivalves, and in particular many unionids, however, presents a unique challenge due to the highly endangered, and thus protected, nature of many of these species and a, generally, limited ability to secure and maintain organisms for testing. The ability to assess effects of toxicant exposure in both laboratory and field studies on unionid reproductive condition and gametogenic development using minimally invasive and nonlethal methods is paramount to the ability to gain information for such species. Typical methods used to assess reproductive development and gametogenesis in bivalves focus on histological analyses, which have most always required organism sacrifice. Recent advances in nonlethal assessment and methodological developments focused on the utilization of nonlethal procedures has, however, provided a new avenue by which evaluation of potential exposure effects might be evaluated. Therefore, this

ii study aimed first to test the ability of a nonlethal gonad biopsy to provide information sufficient for accurate gametogenic assessment of the common unionid, Elliptio complanata.

E. complanata utilized in this study were collected at one of two sites in the Cacapon River of West Virginia (WV). Spring and summer sampling was conducted in order to assess the ability of the biopsy procedure to stage gametogenesis at different points of the reproductive cycle. A biopsy of gonadal fluid was collected from each organism, and the individuals were then sacrificed and preserved for histological analysis in order to make possible a known method of gametogenic staging for comparison and validation of biopsy-based stage determinations. Data obtained using both the histological and biopsy-based approach were compared, and method of biopsy analysis was generated that resulted in a high degree of agreement between histological and biopsy stage determinations. Limitations of the procedure were noted, including a lack of fully representative samples of the various gametogenic cell stages in the biopsy. The influence of this limitation appeared to be more influential to female analysis, however the biopsy procedure was found to predict histological stage determinations with a high degree of accuracy for both sexes of E. complanata, at various stages of development. This procedure was further validated using a similar species, Elliptio insulsa as part of a study on the effects of 17β-estradiol (E2) exposure on gametogenesis and reproductive development in this species.

As a high prevalence of intersex condition has been observed for fish populations residing in the Cacapon River and surrounding river systems, similar effects on reproduction and gamete development were studied for E. complanata in the Cacapon River. The field research sites identified in this study were located in the most upper (Site 1) and lower (Site 2) reaches of the

Cacapon, and were associated with land use activities characterized by mainly agricultural activity

(Site 1) or forest (Site 2). Of primary ecotoxicological interest to Site 1 land use is the high

iii concentration of poultry farming that occurs in this region, and the associated land-application of litter waste that has been shown to potentially contain elevated levels of the primary vertebrate estrogen, E2 and its primary metabolite, estrone (E1), among other potentially toxic xenobiotics.

Rates and timing of application of these wastes, and potential over-application associated with an overabundance of litter in the region, may influence the risk of area surface waters receiving runoff contaminated with these estrogenic compounds and, in turn, might increase the risk of exposure to resident mussels.

Whether vertebrate estrogens play an active role in any reproductive or physiological processes in unionids is currently debated. While a growing body of evidence appears to suggest effects in these organisms following exposure to a variety of estrogenic compounds, including known EDCs, often these exposures are to complex mixtures in the environment, such as sewage treatment plant (STP) effluents that demonstrate some degree of estrogenic activity, while other studies have been criticized for experimental design flaws, a lack of replication, or an absence of quantitative response measures.

Apparent impacts on gamete development were observed, particularly for male E. complanata at Site 1, as a high degree of variance in gamete production was observed for samples collected from this population, particularly in comparison to samples collected from Site 2. In addition, a relocation study on organism growth and gamete development suggests that Site 1 organisms are experiencing growth inhibition and, possibly, increased rates of gamete resorption, though this latter observation could not be statistically verified.

In conjunction with organism sampling at each site, passive sampling devices, known as

Polar Organic Chemical Integrative Samplers (POCIS) were deployed during different periods of the year at each site, and extracts were evaluated for their total estrogenic activity using the

iv recombinant yeast estrogen screen, or YES assay. Results of these tests suggest that the estrogenic character of Cacapon River surface waters is variable, but support the general presence of some chemical(s) that contain inherent estrogenic activity.

In order to further test whether vertebrate estrogens effect unionid gamete development, E. insulsa were exposed in the laboratory to varying doses of E2. Organisms were initially sexed and individuals of each gender were randomly placed into one of four exposure groups. Groups were defined by exposure level; “High Dose” (19 ng/µL E2), “Intermediate Dose”

(1.9 x 10-2 ng/µL E2), “Low Dose” (1.9 x 10-5 ng/µL E2), or solvent control and were injected with

E2 at the start of the experiment, and, again, after five days. A biopsy of gonad was then collected

10-days after the initial dosing event, and again five months after the start of the experiment, after a period of expected active gametogenic development. Organisms were also sacrificed at that time for histological analysis. No significant effects on gamete development were observed for female E. insulsa in any exposure group, however, after the five month period, male E. insulsa were found to exhibit a significant decrease in gamete production in all exposure groups, relative to controls. These effects were observed in histological samples, however similar effects were not detected in analysis of biopsy tissues, though the depressed gamete development among these males is similar to findings for male E. complanata located at Site 1 during field studies, as the same metric was analyzed and a high degree of variance was detected within this sample of the

Site 1 population.

The application of this biopsy method of gametogenic assessment was also utilized for the federally endangered unionid species, Pleurobema clava. Previous surveys had found that one

P. clava population, located in the East Fork West Branch St. Joseph River in Michigan (MI) was exhibiting a lack of recruitment and apparent cessation of reproduction. The population was

v characterized by old, large individuals and no young organisms. Therefore, an investigation was undertaken with the objective to identify the stage, or stages, at which reproduction was failing in this population, in order to better guide future conservation efforts. Additionally, POCIS samplers were deployed seasonally in order to determine the presence or absence of estrogenic compounds in the surface waters proximate to these organisms. An additional study site for P. clava was located in the Allegheny River near Tionesta, Pennsylvania (PA) and served as a reference site for this study, and a surrogate species, Elliptio dilatata was sampled concurrently as sample sizes of P. clava were limited due to the highly endangered status of this species.

Sampling of P. clava in the East Fork indicates recent successful recruitment in this population, as a number of very young organisms were collected and positively identified. However, a high degree of gametogenic variability was observed for males sampled in this population at a time of expected gamete maturity, and no evidence of successful fertilization or glochidial brooding was observed at any time at this site. In addition, while known hosts for P. clava were identified as present at the site, no evidence of glochidial encystment was observed on any fish visually inspected at the site or more closely evaluated in the laboratory.

Based upon findings in this study, the potential for nonlethal sampling of unionids for the assessment of gamete development seems feasible, and was validated for E. complanata in the

Cacapon River. In addition, the procedure developed for E. complanata was tested on E. insulsa as part of an E2 exposure study, and agreement between biopsy-based gametogenic staging and histological staging was strong, supporting the suitability of this methodology for multiple unionid species.

Findings regarding gamete development for E. complanata in the Cacapon River (WV), in light of positive POCIS sampler estrogenicity values detected in several extracts following

vi deployment in the vicinity of these organisms, as well as for E. insulsa tested in the laboratory, fail to refute a potential role for estrogens or estrogenic compounds in the environment in unionid reproduction and gamete development. It is recommended that future studies in freshwater bivalve ecotoxicology attempt to expand the utility of nonlethal methods of reproductive and gametogenic assessment. In addition, further clarification of the possible role estrogens and estrogen mimics in the environment might play in reproductive impairment of unionid mussels is suggested, and the possible significance of xenobiotic exposure to continued unionid declines should remain an area of focus in order to guide ongoing conservation efforts.

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Acknowledgements

I would like to first issue a thanks to, and recognize the contributions of, my advisor, Dr.

Roman Lanno for the critical role he has played with the completion of this research. I am grateful to Roman for his patience, wisdom, advice, and willingness to support my aspirations to pursue research in an area of ecotoxicology that lies somewhat outside the typical focus of his laboratory.

I would also like to say thanks to all of the professionals and friends who have aided me in the successful completion of this work. In particular, thanks to my committee members, Dr. Susan

Fisher, Dr. G. Thomas Watters, and Dr. Linda Weavers, for their efforts, advice, and time invested in this project.

I also want to say a very special thank you to my wife, Megan Sovic. Her patience, love, and support over the course of this research effort have been immense and I very honestly could not have done this without her. From long hikes carrying heavy field equipment to spending days sitting in a cold river recording data and looking for mussels, she never failed to support me and my work, and for that I am eternally grateful. I also have the pleasure of recognizing my daughter,

Julia Marie Sovic, and my second child “to-be” for the drive they have instilled in me to succeed.

I must also recognize and thank my friends and family who have, over the years, provided advice, and shared invaluable knowledge, wisdom, and support. A special thanks to all of those who have assisted in both field and laboratory components of this research, including but in no way limited to, my brother Mike and his wife, Erin Sovic, and my good friends Kody Kuehnl, Adam

Andrews, and Brad Graley for their efforts and assistance in completion of this work.

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Finally, I would like to recognize the contributions and support received from the Robert

H. Edgerley Environmental Toxicology Fund, the Great Lakes Restoration Initiative, and the Helen

M. and Milton O. Lee Fellowship. I would also like to thank the West Virginia Department of

Natural Resources, the Pennsylvania Fish and Boat Commission, the Ohio Department of Natural

Resources, The United States Fish and Wildlife Service, the Columbus Zoo and Aquarium, the OSU

Environmental Science Graduate Program, the OSU Department of Evolution, Ecology, and

Organismal Biology, and the OSU Center for Life Sciences Education, all of which provided tremendous support over the course of my graduate experience at The Ohio State University.

Vita

2002………………………………..Ripley High School

2007………………………………..B.S. Cell and Molecular Biology and Biochemistry, Marshall University

2007 to present……………….Graduate Student, Environmental Science Graduate Program, The Ohio State University

Publications

Krise, K.M., Hwang, A.A., Sovic, D.M., Milosavljevic, B.H. 2011. Macro- and Microscale Rheological Properties of Poly(vinyl alcohol) solutions. Phys Chem B, 115(12): 2759- 2764.

Hurdzan, C.M., Lanno, R.P., Sovic, D.M. 2011. Differential acute toxicity of tetrachlorobenzene isomers to oligochaetes in soil and water: application of the critical body residue concept. Bull Environ Contam Toxicol, 87(3): 209-214.

Sovic, D. M., Lester, L. R., Murray, E. E., Cohenford, M. A. 2008. The Utilization of Bathocuproinedisulfonic Acid as a Reagent for Determining D-glucose and D-galactose Levels in Glycoconjugates. Bioorganic Chemistry, 36(2): 91-95.

FIELDS OF STUDY

Major Field: Environmental Science Graduate Program

Table of Contents

Abstract………………………….……………………….……………………….……………………….………………………..………ii

Acknowledgements. ……….……………………….……………………….……………………….……………………..……..viii

Vita………………………….……………………….……………………….……………………….…………………………….………..x

List of Tables………….……………………….……………………….……………………….…………………………….…………xii

List of Figures………….……………………….……………………….……………………….…………………………….. ..……xv

List of Images………….……………………….……………………….……………………….…………………..…………..……xxi

Chapter 1: An Introduction to Freshwater Mussel Biology and Current Understanding of

Endocrine Disruption in Bivalve Molluscs……………………………………………………..………………..1

Chapter 2: Application of a Nonlethal Biopsy Method for Evaluation of Gametogenesis

in the Freshwater Mussel Elliptio complanata (Bivalvia: Unionidae)……………………………..………6

Chapter 3: Laboratory and field investigations into estrogenic effects on growth

and gametogenesis in freshwater mussels.……………………….……………………..………….……..64

Chapter 4: Investigation into recent observations of recruitment failure in the East

Fork West Branch St. Joseph River population of the federally endangered

freshwater mussel, Pleurobema clava (Bivalvia: Unionidae)……………………..……..……….139

Chapter 5: Conclusion………….……………………….……………………….……………………….………….………..…178

References………….……………………….……………………….……………………….………………………….……….…..183

LIST OF TABLES

Table 2.1 – Biopsy-Generated Spermatogenic Cell Classifications in male E. complanata……….…29

Table 2.2 – Biopsy-Generated Gametogenesis Classifications in male E. complanata...... 30

Table 2.3 – Biopsy-generated spermatogenic cell classifications in male E. complanata……………31

Table 2.4 – Cohen’s Kappa Statistic Strength of Agreement Scale……………………………………………..32

Table 2.5 – Gametogenesis classifications in male E. complanata, modified from Barber

(1996) and Yokely (1972)………………………………………………………………………………………………33

Table 2.6 – Threshold value approximations for male E. complanata used in the

determination of area-based histological analysis………………………………………………………..42

Table 2.7 - Biopsy-Generated Gametogenesis Classifications in male E. complanata………………..52

Table 2.8 – Histology-based gametogenic stage classification for female E. complanata……….….53

Table 2.9 – Back-transformed mean and 95% Confidence Values for ova measured in

histological and biopsy samples from female E. complanata according to

gametogenic stage assigned………………………………………………………………………………………...58

Table 2.10 – Threshold values determined for biopsy-based identification of

gametogenic stage in female E. complanata based on ova areas observed in

collected biopsy fluid…………………………………………………………………………………………….……..62

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Table 3.1 – YES assay row compositions of POCIS sampler extracts assayed. % POCIS

represents the maximum percent of the total extracted sorbent mass assayed

during each test……………………………………………………………………………………………………….…103

Table 3.2 – POCIS extract characteristics for YES assay testing………………………………………………..104

Table 3.3 – Site/Sex differences observed in morphometric measures and associated

p-values resulting from two-sample Student’s T-test; all α = 0.05………………………………109

Table 3.4 – Observed gender differences for E. complanata and resulting statistics

following Chi-Square test (α = 0.05)………………………………………………………………….………..110

Table 3.5 – EC50 and EC20 values and associated 95% confidence intervals for 17β-estradiol

standard dilution series curves from each POCIS extract YES assay microplate……….….119

Table 3.6 – Maximal absorbance values observed for each POCIS composite extract and

corresponding E2 Concentrations (M) from standard dilution series analyses……………120

Table 3.7 – Mean differences in female AAF values observed for acini within E. insulsa

exposed to the three levels of 17β-estradiol versus solvent control…………………………..127

Table 4.1 – Deployment periods for POCIS samplers at both EF and AL study sites…………………160

Table 4.2 – Gonad viability classifications for P. clava sampled in 2013 from the EF and

AL sites……………………………………………………………………………………………………………………….166

Table 4.3 – Gonad viability classifications for E. dilatata sampled in 2013 from the EF and

AL sites……………………………………………………………………………………………………………………….167

Table 4.4 – POCIS estrogenicity values as determined by the YES assay…………………………………..176

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Table 4.5 – Viability (%) of mature glochidia collected from AL P. clava (n = 2) in early

July, 2012……………………………………………………………………………………………………………………177

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LIST OF FIGURES

Figure 1.1 The unionid life cycle………………………………………………………………………………………………….5

Figure 2.1 Average percentage values for proportion of atretic tissue observed in

histological sections of gonad tissue of male E. complanata………………………………………..35

Figure 2.2 Average percentage values for proportion of atypical tissue observed in

histological sections of gonad tissue of male E. complanata………………………………………..36

Figure 2.3 – Histology-based freshwater mussel male gametogenic classification protocol

generated utilizing E. complanata from the Cacapon River, WV…………………………………..43

Figure 2.4 – Gametogenic stage observed for male E. complanata in Spring (May)

2013 and Summer (late July/early August) 2013…………………………………………………………..44

Figure 2.5 - Male E. complanata gametogenic stage classifications as generated by the

area-based and transect-based histological approaches (total nSpring + Summer = 21)…………45

Figure 2.6 – Individual value plot indicating threshold value for atretic and non-atretic

categorization in male biopsy analysis (n = 21 organisms sampled)………………………………47

Figure 2.7 - Individual value plot indicating threshold value for Typical and Atypical

categorization in male biopsy analysis (n = 21)…………………………………………………………….48

Figure 2.8 - Biopsy-based male freshwater mussel gametogenic classification protocol

generated utilizing E. complanata from the Cacapon River, WV…………………………………..49

Figure 2.9 - Male E. complanata gametogenic stage classifications as generated by

the area-based histological method and biopsy-based analysis

(total n Spring + Summer = 21)……………………………………………………………………………………………….50

Figure 2.10 - Male E. complanata gametogenic stage classifications as generated by

the transect-based histological method and biopsy-based analysis

(total n Spring + Summer = 21)……………………………………………………………………………………………….51

Figure 2.11 – Gametogenic stage observed for female E. complanata in Spring (May) 2013

and Summer (late July/early August) 2013 based upon histological analysis……………….57

Figure 2.12 – Ova area distributions (untransformed) for developing and mature ova

observed in E. complanata samples according to measurement method……………………..60

Figure 2.13 – Untransformed ova area distributions resulting from histological and

biopsy-based analysis for female E. complanata classified as Spent/Resorbing…………….61

Figure 2.14 – Female E. complanata gametogenic stage classifications as generated by

histological and biopsy-based analysis………………………………………………………………………….63

Figure 3.1 – Gametogenic stages initially identified following visual inspection of

histological samples for male (nSite 1 = 5, nSite 2 = 7; left) and female (nSite 1 = 14,

nSite 2 = 12; right) E. complanata from two sites in the Cacapon River, WV………………….106

Figure 3.2 – Initial mass (g) distributions for E. complanata from each of the two

study sites………………………………………………………………………………………………………………….107

Figure 3.3 - – Initial length (mm) distributions for E. complanata from each of the

two study sites……………………………………………………………………………………………………………108

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Figure 3.4 – Observed gender ratios as determined by biopsy analysis and expected

counts for E. complanata at Sites 1 and 2 (nsite 1 = 85; nsite 2 = 102)……………………………..111

Figure 3.5 – Gametogenic stages observed for E. complanata of both sexes at the

two Cacapon River study sites in May 2013; for each site, n = 5/sex…………………………..112

Figure 3.6 - Gametogenic stages observed for E. complanata of both sexes at the

two Cacapon River study sites in late July/early August 2013 (Site 1 nmale = 6,

nfemale = 3. Site 2 nmale = 6, nfemale = 4)…………………………………………………………………………..113

Figure 3.7 – Average AAF values calculated for E. complanata at each site included in

this study (Males nspring = 10, n summer = 12; Females nspring = 10, nsummer = 7)…………………114

Figure 3.8 – Average GAF values calculated for E. complanata at each site included in

this study (Males nspring = 10, n summer = 11; Females nspring = 10, nsummer = 7); one

outlier value was omitted for Males, Site 1 from the Summer sampling…………………….115

Figure 3.9 – Average 20-month mass changes (g) for control (R1 and R2) and relocated

(Exp1 and Exp2) E. complanata (nR1 = 16, nR2 = 14, nExp1 = 11, nExp2 = 13) in the

Site 1/Site 2 relocation study……………………………………………………………………………………..116

Figure 3.10 – Tukey pairwise comparisons for differences of means for 20-month mass

changes (g) in E. complanata relocated between Site 1 and Site 2 (nR1 = 16,

nR2 = 14, nExp1 = 11, nExp2 = 13)……………………………………………………………………………………..117

Figure 3.11 – 17β-estradiol standard (positive control) response data generated from

POCIS extract analyses; each point represents E2 standard response data from

each of six test plates…………………………………………………………………………………………………118

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Figure 3.12 - E2 concentrations (M) corresponding to average maximal responses

(corrected absorbance) of POCIS extracts; bars indicate 95% confidence

intervals and Site 2 June 2012 data point represents upper limit of 99% CI for

negative control, as absorbance values were below LOD……………………………………………121

Figure 3.13 – Median, Inter-quartile range, and total range of observed ova areas (µm2)

from biopsy samples collected at time = 10 days following initial dosing of

organisms (n = 5/group)……………………………………………………………………………………………..126

Figure 3.14 – Average AAF values observed for E2 exposed E. insulsa versus controls; bars

represent 95% confidence intervals……………………………………………………………………………128

Figure 3.15 – Median, Inter-quartile range, and total range of observed ova areas (µm2)

from biopsy samples collected at time = 5 months following initial E2 dosing

of E. insulsa (n = 5/group, Control and High dose; n = 4/group Low and

Intermediate dose) with E2………………………………………………………………………………………..133

Figure 3.16 – 95% confidence intervals for the difference of E2 exposure level mean and

control mean % atretic values (Dunnett Multiple Comparisons with a Control test)

for female E. insulsa……………………………………………………………………………………………………134

Figure 3.17 – Gametogenic stage determinations for male E. insulsa sampled 5-months

post initiation of E2 exposure study (n = 20) using the biopsy-based approach as

well as the histological approach (see Figures 2.3 and 2.8 for methodologies)…………..135

xviii

Figure 3.18 – Dunnett Simultaneous 95% CIs. Intervals represent 95% confidence intervals

for the difference between 5-month post-exposure mean AAF values among

Control organisms (n = 5) and mean AAF values among organisms at each

exposure level (n = 5 organisms per exposure level)…………………………………………………..136

Figure 3.19 – Boxplots of average proportion (%) of atretic tissue observed within

measured acini in E. insulsa (n = 5/group) 5-months post E2 exposure………………………137

Figure 3.20 - Boxplots of average proportion (%) of atretic tissue observed in biopsy

samples analyzed from E. insulsa (n = 5/group) 5-months post E2 exposure………………138

Figure 4.1 – Total mass (g) and maximum valve length (mm) distributions for P. clava

at EF……………………………………………………………………………………………………………………………162

Figure 4.2 – Total mass (g) and maximum valve length (mm) distributions for P. clava

at AL…………………………………………………………………………………………………………………………..163

Figure 4.3 - Total mass (g) and maximum valve length (mm) distributions for E. dilatata

at EF……………………………………………………………………………………………………………………………164

Figure 4.4 - Total mass (g) and maximum valve length (mm) distributions for E. dilatata

at AL…………………………………………………………………………………………………………………………..165

Figure 4.5 – Gametogenic stages observed in EF P. clava collected and sampled for a

biopsy of gonadal fluid in 2013…………………………………………………………………………………..168

Figure 4.6 – Gametogenic stages observed in AL P. clava collected and sampled for a

biopsy of gonadal fluid in 2013…………………………………………………………………………………..169

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Figure 4.7 - Gametogenic stages observed in EF E. dilatata collected and sampled for a

biopsy of gonadal fluid in 2013…………………………………………………………………………………..170

Figure 4.8 - Gametogenic stages observed in AL E. dilatata collected and sampled for a

biopsy of gonadal fluid in 2013…………………………………………………………………………………..171

Figure 4.9 – Expected and observed number of P. clava of each sex positively identified

at EF (total n = 21)………………………………………………………………………………………………………172

Figure 4.10 – Expected and observed number of P. clava of each sex positively identified

at AL…………………………………………………………………………………………………………………………..173

Figure 4.11 – Expected and observed number of E. dilatata of each sex positively

identified at EF……………………………………………………………………………………………………………174

Figure 4.12 – Expected and observed number of E. dilatata of each sex positively

identified at AL…………………………………………………………………………………………………………..175

LIST OF IMAGES

Image 2.1 – Acinus (40x objective) representing Spent/Resorbing gametogenic classification

in male E. complanata………………………………………………………………………………………………….34

Image 2.2 – Acinus (40x objective) representing Early Atypical gametogenic classification

in male E. complanata………………………………………………………………………………………………….37

Image 2.3 – Acinus (40x objective) representing Late Atypical gametogenic classification

in male E. complanata………………………………………………………………………………………………….38

Image 2.4 – Acinus (40x objective) representing Early Active gametogenic classification in

male E. complanata………………………………………………………………………………………………………39

Image 2.5 - Acinus (40x objective) representing Late Active gametogenic classification in

male E. complanata………………………………………………………………………………………………………40

Image 2.6 - Acinus (40x objective) representing Mature gametogenic classification in male

E. complanata………………………………………………………………………………………………………………41

Image 2.7 – Biopsy (40x objective) of male gonad from E. complanata showing mature

spermatozoa (SZ), spermatocytes (SC), spermatids (SP) and an atypical pathway

morulae (SM)………………………………………………………………………………………………………………………46

Image 2.8 – Acinus (40x objective) representing Mature gametogenic classification in

female E. complanata……………………………………………………………………………… …………………54

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Image 2.9 – Acinus (40x objective) representing Developing gametogenic classification in

female E. complanata…………………………………………………………………………………………………..55

Image 2.10 – Acinus (40x objective) representing Atretic gametogenic classification in

female E. complanata…………………………………………………………………………………………………..56

Image 2.11 – Biopsy of female gonad (4x objective) from E. complanata showing mature (M),

developing (D) and atretic (A) ova……………………………………………………………………………………….59

Image 3.1 – Aereal photograph of the Cacapon River at Wardensville, WV (modified

from Constantz et al, 2003. Portrait of a River – The Ecological Baseline of the

Cacapon River.) Site 1 is located in the bottom-left of the image, just upstream of

the town’s sewage settling ponds.……………………………………………………………………………….99

Image 3.2 – Photograph taken looking upstream in the Cacapon River at Site 1. All organisms

were sampled along the left descending bank in the area just above the

riffle seen in this image……………………………………………………………………………………………..100

Image 3.3 – Aereal photograph of the Cacapon River near Great Cacapon, WV

(modified from Constantz et al, 2003. Portrait of a River – The Ecological Baseline

of the Cacapon River.) Site 2 is located near the center of the image, as the river

winds toward the mouth. ………………………………………………………………………………..……....101

Image 3.4 - Photograph taken looking downstream from a low-water bridge

crossing the Cacapon River at Site 2. All organisms were sampled near midstream

just below the riffle seen in this image………………………………………….…………………………..102

Image 3.5 – Raceways located at the Freshwater Mussel Conservation and Research

Center in Shawnee Hills, OH……………………………………………………………………………………….105

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Image 3.6 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post

initial exposure to 17β-estradiol at the lowest concentration tested

(1.9 x 10-5 ng/µL)………………………………………………………………………………………………………..122

Image 3.7 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post

initial exposure to 17β-estradiol at the intermediate concentration tested

(1.9 x 10-2 ng/µL)………………………………………………………………………………………………………..123

Image 3.8 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post

initial exposure to control solvent (27% EtOH)……………………………………………………………124

Image 3.9 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post

initial exposure to 17β-estradiol at the highest concentration tested (19 ng/µL)……….125

Image 3.10 – Stained portion of female E. insulsa biopsy tissue collected 5 months post

initial exposure to control solvent (27% EtOH)……………………………………………………………129

Image 3.11 – Stained portion of female E. insulsa biopsy tissue collected 5 months post

initial exposure to 17β-estradiol at the lowest concentration tested

(1.9 x 10-5 ng/µL)………………………………………………………………………………………………………..130

Image 3.12 – Stained portion of female E. insulsa biopsy tissue collected 5 months post

initial exposure to 17β-estradiol at the intermediate concentration tested

(1.9 x 10-2 ng/µL)………………………………………………………………………………………………………..131

Image 3.13 – Stained portion of female E. insulsa biopsy tissue collected 5 months post

initial exposure to 17β-estradiol at the highest concentration tested (19 ng/µL)……….132

xxiii

Image 4.1 – Examples representing the size range observed for P. clava in the East Fork

West Branch St. Joseph River……………………………………………………...... …………………….…161

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Chapter 1: An Introduction to Freshwater Mussel Biology and Current Understanding of

Endocrine Disruption in Bivalve Molluscs

The group of organisms collectively referred to as pearly or freshwater mussels includes the bivalve molluscs of the order Unionioda. Of all taxa classified within this order, the unionids

(family Unionidae) are distributed worldwide, comprise approximately 677 named species in total

(McElwain and Bullard, 2014), and, while highly speciose, are considered one of the world’s most imperiled faunal groups (Haag and Williams, 2014; Williams et al., 1993). The impacted nature of this group has resulted in significant effort on behalf of the scientific community to better understand both the unique biology of these organisms, as well as the potential factors contributing to their observed declines.

Freshwater mussels employ a unique and relatively complex life cycle, as is depicted in

Figure 1.1 (Watters, 2007). It is generally accepted that two differing reproductive strategies exist within the unionids, the differentiating factor between these approaches being the duration of glochidial brooding. However, the general life cycle of all unionids, while complex, appears to be, essentially, the same. While a few hermaphroditic and “occasionally hermaphroditic” species have been observed (Van Der Schalie, 1970; Heard, 1979; Kat, 1983), unionid species are typically dioecious, with adults developing either ova or sperm in diffuse gonadal tissues located throughout the visceral mass. Though males release sperm freely into the water column, often packaged by the thousands into specialized structures referred to as, “sperm balls” or spermatozeugmata, fertilization occurs internally. Females siphon male gametes from the water and fertilization of ova takes place within the suprabranchial chamber of the gill. Fertilized ova

1 are then moved to specialized regions of the gill (marsupia) and develop into an obligate parasitic life stage known as a glochidium, which requires an encystment period on a host organism, usually a fish, prior to transformation to the juvenile stage. Full elucidation of this component of the unionid approach to reproduction, development, and maturation was not realized until the late

1800s, when experiments with Anodonta confirmed the notion that glochidia were, in fact, a parasitic larval life stage of the mussel and not a separate parasitic species, nor a typical larval stage of development, as had been previously proposed (Watters, 2007). Following successful encystment and transformation on the appropriate host, the juvenile mussel drops to the substrate and continues growth and development to maturity.

The unique and highly complex life cycle employed by freshwater mussels, as well as the general biology of these organisms, opens the door to potential modulation and ultimate disruption of reproduction by exogenous chemicals and xenobiotics found in their environment.

Since adult organisms are filter feeders, the risk of uptake and exposure to xenobiotics can be high, both for water-borne contaminants, as well as those sorbed to suspended materials. In a field study of pharmaceutical bioaccumulation across multiple trophic levels, elevated concentrations (defined as > 130 µg/kg) of both the antidepressant sertraline, as well as its primary metabolite, desmethylsertraline, were observed in tissues of both the Asian clam,

Corbicula fluminea, as well as two unionid species, the pondhorn, Uniomerus tetralasmus, and the paper pondshell, Utterbackia imbecillis (Du et al., 2014). In addition, recent research found an unusually high sensitivity of glochidial and juvenile life stages of some unionid species to a number of environmental contaminants (Bringolf et al., 2007a; Bringolf et al., 2007b; Bringolf et al., 2007c; Newton and Bartsch, 2007; Wang et al., 2008; Strayer and Malcom, 2012). However, due to the relative lack of sufficient knowledge regarding many of the specific biochemical and physiological processes that occur in unionids, as well as the limited amount of ecotoxicological

2 research that has been conducted utilizing these organisms, the ultimate risk of exposure effects of many chemicals to these organisms remains unclear. There may be no better current example of this than when considering the vertebrate hormones, including 17β-estradiol (E2) and its xenoestrogen mimics, and the potential effects these compounds may exert on unionid reproduction and development. These and a variety of other known compounds have now been long identified as endocrine disrupting chemicals (EDCs) that have the potential to exert adverse effects across a diverse group of taxa following exposure, particularly among aquatic species

(Sumpter, 2005). Estrogens and xenoestrogens have been shown to elicit adverse effects on an array of aquatic species following exposure to these compounds at environmentally relevant concentrations (Janer and Porte, 2007; Frye et al., 2013). Studies have also suggested adverse effects on a number of bivalve species, including a limited number of studies utilizing unionids

(Leonard et al., 2014a; Loenard et al., 2014b; Gagné et al., 2001; Jobling et al., 2004; Keller et al.,

2007). Studies to date, however, have failed to fully establish a functional role for vertebrate or vertebrate-like hormones such as E2 in bivalves, have generally failed to detect the presence of an estrogen receptor similar to that commonly found in vertebrates, have been variable in the detection of adverse effects following estrogen and xenoestrogen exposure, and have been criticized for a number of issues related to experimental design (Scott, 2012; Scott, 2013). That said, data generated by a number of studies on a various bivalve species have either supported the hypothesis that vertebrate or vertebrate-like steroid hormones play a role in bivalve reproduction, or have suggested various effects associated with exposure to individual estrogens and xenoestrogens, as well as complex environmental mixtures, such as from sewage treatment or other industrial effluents (Ciocan et al., 2011; Croll and Wang, 2007; Leonard et al., 2014a;

Loenard et al., 2014b; Liu et al., 2008; Gagné et al., 2001; Jobling et al., 2004; Keller et al., 2007).

The stark contrast that currently exists between those studies supporting and those refuting the

3 hypotheses of either a physiological role for estrogens in bivalves or effects related to estrogen and/or xenoestrogen exposure in bivalves suggests a gap in understanding, and highlights the need for additional research to begin to clarify the debate.

This complex topic of endocrine disruption in freshwater mussels is addressed in the work described following this introduction. First, in Chapter 2, a nonlethal method for the evaluation of gametogenesis in these organisms is presented. This chapter details the design and validation of a quantitative approach to assessing both histological and biopsy samples of unionid reproductive tissue. The application of this method is then highlighted in Chapters 3 and 4, which describe field and laboratory studies that were conducted, and that involved multiple unionids, including three Elliptio species (E. complanata, E. insulsa and E. dilatata), as well as the federally endangered clubshell, Pleurobema clava. These field studies investigated the reproductive condition and seasonal reproductive patterns of unionids found in a number of locations, including the Cacapon River in West Virginia, the East Fork West Branch St. Joseph River in

Michigan, and the Allegheny River in Pennsylvania. An additional component of this research that is presented involved the deployment of passive sampling devices used to test for the presence or absence of estrogenic compounds in surface waters associated with the unionid populations sampled. Chapter 3 also describes an experiment carried out in the laboratory that tests the effects of 17β-estradiol exposure on gametogenesis in both male and female adult E. insulsa.

Finally, a concluding discussion of the findings of this research is provided in Chapter 5, along with suggestions for future research efforts in both unionid ecotoxicology and conservation biology.

Figure 1.1 – The unionid life cycle (from Watters, 2007).

Chapter 2: Application of a Nonlethal Biopsy Method for Evaluation of Gametogenesis in the

Freshwater Mussel Elliptio complanata (Bivalvia: Unionidae)

Introduction

Nearly 300 species and subspecies of freshwater mussels have been known to occur in

North America (Williams et al., 1993), with nearly two-thirds of these species considered to be of some degree of conservation concern (Haag and Williams, 2014). These organisms are considered one of North America’s most endangered faunal groups (Augspurger et al., 2007; Christian and

Harris, 2008; Haag and Williams, 2014). Traditionally, taxonomic classification of these organisms has been based on soft part anatomy and shell morphometry (Bogan and Roe, 2008), though recent advances in the application of genomic data has led to classification re-evaluation based upon modern phylogenetics (2008) and will likely result in greater species diversity than previously assumed (Haag and Williams, 2014). At the time of the publication of Conservation

Status of Freshwater Mussels of the United States and Canada, Williams et al. (1993) identified approximately 72% of these organisms as extinct, endangered, threatened, or of special concern.

That statistic, while alarming, appears to have only grown in recent years, as approximately 75% of North American species are now considered extinct or to some degree imperiled (Haag and

Williams, 2014). This number may likely continue to grow as reclassifications resulting from ongoing phylogenetic analyses are made.

It is clear that declines in freshwater mussel populations have been widely documented, particularly in recent decades, and future extinction rates for North American freshwater mussel

6 fauna have been estimated at more than six percent per decade (Ricciardi and Rasmussen, 1999).

While in some instances population effects may be clearly associated with known disturbances, such as stream fragmentation, flow regulation, channelization, dredging, sediment loading, and discharge of toxic contaminants from municipal and industrial sources (1999), in many cases, the specific cause(s) of observed population declines remain uncertain, thus complicating conservation efforts (Haag and Williams, 2014). Such efforts are further complicated when the specific biology of a target species is largely unknown.

Investigations into health, gametogenesis, spawning period, and reproductive condition of adult freshwater mussels and other molluscs, as well as their utilization in toxicity testing, have historically required organism sacrifice, as the primary methods employed in these investigations have relied on the histological analysis of gonadal tissues (Van der Schalie & Van der Schalie, 1963;

Yokely, 1972; Smith, 1978; Zale and Neves, 1982; Smith 1988; Gordon and Smith, 1990; Jirka and

Neves, 1992; Amyot and Downing, 1998; Grande et al., 2001; Smith, et al., 2003; Henley et al.,

2007). In many cases, such analyses are used to evaluate endpoints of gamete production or reproductive activity, whether for the elucidation of reproductive timing within a population

(Smith et al. 2003), evaluation of reproductive health (Watermann et al., 2008), or as a potential response to toxicant exposure (Ortiz-Zarragoitia and Cajaraville, 2006). Endpoint measures commonly employed in these histological investigations include average ova diameter/area, sex ratios, incidence rates of intersex condition, gametogenic stage determination, and calculation of various gonad and condition indices (Cek and Sereflisan, 2011; Haggerty et al., 1995; Sereflisan,

Cek, and Sereflisan, 2013; Weaver, Pardue, and Neves, 1991; Yeager and Neves, 1986).

An obvious barrier to obtaining important information about already threatened and endangered mussel species is apparent, as the use of lethal means to generate necessary data may further impact already imperiled populations. The development and utilization of nonlethal

7 methods for the evaluation of general and reproductive health in freshwater mussels may provide a route by which this barrier can be circumvented.

Recent studies have demonstrated the utility of nonlethal biopsy techniques for various unionid tissues, including mantle (Berg et al., 1995), foot (Naimo et al., 1998), hemolymph

(Gustafson, 2005) and gonad (Saha and Layzer, 2008). Such nonlethal techniques may provide a more conservation-conscious approach to unionid research and a means to more effectively study threatened and endangered unionid species. Endpoint measures employed via the various nonlethal biopsy techniques that have been published for unionid mussels include glycogen content of foot tissue (Naimo et al., 1998), DNA analyses (Karlsson et al., 2013), metabolic profile characterization (Roznere et al., 2014) and determination of sex in non-sexually dimorphic species

(Saha and Layzer, 2008). The work by Saha and Layzer (2008) has particular application to the present study, as this procedure has been shown to consistently collect enough developing gonadal tissue to make accurate gender determinations in multiple non-sexually dimorphic freshwater mussel species, with no observable effect on either short- or long-term mortality of organisms sampled. In addition, method validation procedures included in that study suggest that, at minimum, a simplified gametogenic stage classification scheme aligns well with information collected from histological analyses (2008). Henley (2002) concluded that this type of nonlethal biopsy had the potential to allow for gametogenic stage determinations in the absence of organism sacrifice for two unionid species (Utterbackia imbecillis and Villosa iris), though the need for further testing on additional unionid species was suggested.

This study addresses the question of whether collection and analysis of a nonlethal biopsy of developing gonad tissue can serve as a viable alternative to traditional histological examination of the viscera for the assessment of gametogenesis in the Eastern Elliptio, E. complanata

(Lightfoot, 1786). Specifically, this study aims to test the following hypothesis:

1) If the gonad in E. complanata is diffusely distributed throughout the visceral mass and

a nonlethal biopsy of gonadal tissue in E. complanata consistently dislodges and

collects a fully representative sample of the developing gonadal tissue, then analysis

of the contents of that biopsy should consistently predict the gametogenic stage

designated using the traditional histological approach.

The primary objective of this study is to compare gametogenic stage classifications generated from the analysis of both biopsy samples and histological preparations of those biopsied individuals. These comparisons are completed for individuals collected from two separate sites in the Cacapon River in West Virginia at different points within the reproductive cycle.

Materials and Methods

Site Selection and Mussel Collection

E. complanata individuals were collected in May 2013 and late July 2013 from two sites in the Cacapon River, West Virginia. Site 1 was located at approximately 39° 05’ 31.1” N, 78° 35’

42.9” W (elevation 291 m) in Wardensville, Hardy County, West Virginia (WV), while Site 2 was located at approximately 39°36’26.31” N, 78°17’29.21” W (elevation 133 m), just downstream of a low-water bridge on PowerHouse Road (CR 9/12) near Berkeley Springs, Morgan County, WV.

During each sampling event, 10 individuals were collected via hand-grab with aid of a glass- bottom bucket from each of the two Cacapon River sites. Organisms at Site 1 were found immediately off of the left descending bank in a shallow run exhibiting a high density of E. complanata. Organisms were observed more sporadically at Site 2, though most were collected from a mid-stream run. All individuals collected were weighed, biopsied, and sacrificed for histological analysis. In order to preserve tissues for analysis, both the biopsied tissue and the

9 visceral mass of each sacrificed individual were immediately fixed in 10% Neutral Buffered

Formalin (NBF) at the site.

Biopsy Procedure and Tissue Collection

The nonlethal gonad biopsy procedure was carried out according to the description in

Saha and Layzer (2008), but adapted for utility in the field. Prior to sampling, syringes (Excel 3 cc) equipped with 18-gauge needles were preloaded with 0.5 mL deionized water and recapped. At the site, organisms were located and valves were gently pried apart using snap ring pliers and held open by inserting a syringe cap between the valves. The needle of a preloaded syringe was then inserted through the epithelial wall of the foot at an area slightly anterior to the mid-point of the foot/visceral mass complex and into the region of the gonad within the visceral mass.

Approximately 0.5 mL deionized water was injected into the visceral mass in order to dislodge developing gametes, and 0.20 - 0.25 mL of the resulting gonadal fluid was extracted back into the syringe. This fluid biopsy containing fresh gametes was then expelled into a labeled microcentrifuge tube (1.5 mL) prefilled with 750 µL NBF in order to preserve collected gametes for later analysis. Following the biopsy, each organism was immediately sacrificed by separation of both anterior and posterior adductors and excision of the foot/visceral mass complex and immersion in a pre-labeled 50-mL centrifuge tube filled with 40 mL NBF.

Histological Processing and Analysis

Following biopsy collection in the field, organisms were sacrificed, viscera excised, and tissues fixed in NBF for future processing. In the laboratory, a frontal section (approx. 0.5 mm) was cut from each visceral mass at the approximate midpoint of the anterior/posterior length.

Samples were dehydrated in alcohol and paraffin embedded prior to sectioning and staging, and were stained with a standard HE (hematoxylin and eosin) stain. Five acini from each of two

10 sections were imaged and analyzed using Motic Images Plus 2.0 software. In order to select acini for this analysis, a portion of one histological section was imaged at 4x and a 3 x 3 rectangular grid

(approx. 750 µm x 1000 µm per field) was overlaid across the image. The outermost acinus located within each of the four corner fields of the grid was selected for measurement, in addition to the centermost complete acinus. In the case that 5 complete acini could not be located, the sample was re-imaged after randomly moving the microscope stage along both X- and Y- axes. This process was repeated for each of the two sections examined.

For male samples, two approaches to histological analysis of the gonadal tissue were employed and compared. First, average areas (µm2) occupied by each of four tissue categories –

(1) developing cells (spermatocytes and spermatids), (2) mature cells (spermatozoa), (3) atretic tissue and cellular debris, and (4) atypical spermatogenic tissue (sperm morulae, sertoli cells, and amorphous inclusions) – within each of the 10 acini analyzed per organism, were measured (40x objective) using MOTIC Images Plus 2.0 ML imaging software, and a proportion of each identified cell type to the total area measured was calculated. Stages of development for individual spermatogenic cells observed were made according to cell morphology and staining characteristics (Table 2.1), and were based on previous descriptions (Cek and Sereflisan, 2011;

McElwain and Bullard, 2014). An estimate of the numbers of developmental and mature cells represented in the total area measured per sample from the area-based method was then calculated as the sum total of the product of the total measured area occupied in each acinus by each of the two cell stages (µm2) and the average number of cells observed per µm2 for each respective cell stage (see Equation 1).

퐸푞푢푎푡푖표푛 1: 푥푐 = ∑(퐴푐,푖 × 푎푐) 푖=1

Where 푥푐 is the estimated number of cells observed per stage, c is cell stage

(developmental/mature), levels of 푖 are individual acini measured per organism, 퐴 is the measured area (µm2) for each cell stage per acinus, and 푎 is the average number of cells of each stage (c) per unit measure (µm2). Estimated cell counts generated from the first histological approach for mature and developmental cells, in addition to proportions of atretic and atypical tissues observed, were used in development of staging criteria and for comparisons with cell count-based proportions generated by the alternative, transect-based, approach, as well as with a biopsy-based method, described later. For the alternative histological approach, a single transect was drawn along the greatest length of each imaged acinus, and all cells contacting this transect were identified and enumerated. Cell counts were used to calculate proportions for each identified cell type, similar to those calculated in the area-based approach, and these data were used to make gametogenic stage determinations, according to classifications modified from

Barber (1996) and Yokely (1972).

For females, gametogenic stages were histologically determined based upon proportion of the gonad occupied by either atretic, developmental or mature tissue. In order to select acini for this analysis, as was described earlier, a portion of one histological section was imaged at 4x and a 3 x 3 rectangular grid (approx. 750 µm x 1000 µm per field) was overlaid across the image.

The outermost acinus located within each of the four corner fields of the grid was selected for measurement, in addition to the centermost complete acinus. In the case that 5 complete acini could not be located, the sample was re-imaged after randomly moving the microscope stage along both X- and Y- axes. Areas (µm2) occupied by each of the three tissue types (atretic, developmental, mature) were measured for each acinus selected for analysis and relative proportions were used in the development of histological staging criteria. This process was conducted twice for a total analysis of 10 acini per organism.

Stages of development for developing and mature ova observed were made according to cell morphology and staining characteristics, and were simplified from descriptions provided in

Cek and Sereflisan (2011) into a two-category classification scheme (Developing Oocytes/Mature

Oocytes) (Table 2.2). Acini were selected at random for measurement of ova by identifying the center-most complete acinus after moving the microscope stage randomly along both the X- and

Y- axes under 4x magnification. Developing and Mature ova within the acinus were identified and areas (µm2) were recorded. The irregular, outer vitelline membrane and albuminous space surrounding vitellogenic ova were not included in the measured areas, and only those ova for which the histological section passed through the nucleus were included in the analysis. This procedure was repeated until a total of 30 ova areas had been obtained or until the stage had been repositioned 25 times with no additional ova observed. Measured areas were Square Root transformed in order to meet the parametric statistics assumption of normality and the two- sample Student’s t-test was utilized to compare area distributions among Mature individuals with those classified Developmental according to histological analysis.

Biopsy Preparation

In the laboratory, each fixed, fluid biopsy was concentrated, stained, and imaged for analysis of collected tissues. Samples were effectively pelleted by centrifugation for 5 minutes at high speed (> 192 g). NBF solution was decanted and samples were then treated with a sequence of staining solutions (one to two minutes per solution), each followed by a distilled (DI) water wash and repeated centrifugation for 5 min. The staining sequence was as follows: 25 µL hematoxylin, 100 µL 1 mg/mL sodium bicarbonate solution, 100 µL DI water, 25 µL eosin, 1000 µL

DI water. Following this sequence, the final supernatant was pipetted down to 100 µL total volume, the pellet was resuspended and, if necessary for microscopic analysis, diluted (1:10 –

1:50) with distilled water, and a 10-µL aliquot was transferred to a clean glass slide for initial sex determination.

Male Biopsy Tissue Analysis

For male E. complanata, a 10-µL aliquot of prepared biopsy was transferred to a Neubauer hemocytometer, imaged (40x objective), and analyzed using Motic Images Plus 2.0 ML software.

All gametes observed within, at minimum, four outer 0.0625 mm2 squares were imaged, identified, and enumerated. For those samples exhibiting low total cell counts, additional fields were included in the analysis until at least one developmental or mature cell stage had been observed a minimum of 25 times. Stages of development for individual spermatogenic cells observed were made according to cell morphology and staining characteristics (Table 2.3), and were based on descriptions provided in Cek and Sereflisan (2011). Proportions of each cell type identified were calculated as percentages of the total cells observed from the biopsy. These data, paired with gametogenic stage information collected through the histological approach previously described, were used to generate biopsy-based gametogenic stage classifications.

Female Biopsy Tissue Analysis

For female E. complanata, a 10-µL aliquot of prepared biopsy was transferred to a clean glass microscope slide, and was viewed directly at 4x for analysis of developing ova. Ova present in the biopsy sample were imaged and analyzed using Motic Images Plus 2.0 software. Areas (µm2) of all imaged ova were measured, recorded, and subject to square root transformation to meet the assumption of normality for parametric statistical analysis. For those samples exhibiting low total cell counts, additional aliquots were included in the analysis until at least 25 ova had been observed and measured, or until the biopsy had been exhausted. Transformed data were analyzed using the Two-sample Student’s t-test to compare biopsy data among histologically

14 classified Mature individuals with that among those classified as Developmental. Oocyte areas recorded from biopsy sampling were further compared with histological data and threshold area values were determined to classify each measured ovum according to one of three possible developmental classifications; (1) Developing (2) Mature or (3) Resorbing/Atretic.

Staging Criteria, Statistical Analyses and Protocol Comparisons

Initial gametogenesis stage classifications were modified from Barber (1996) and were generated using results of the area-based histological analyses and estimated cell counts in order to obtain a more quantitative classification scheme for gametogenesis in E. complanata. For males, these classification criteria were then utilized to stage samples by the transect-based approach, in order to compare reliability of this alternative histological method of analysis.

Untransformed male biopsy-generated proportions of each measured cell stage/type among all acini measured were both visually (individual value plots, scatterplots) and quantitatively (mean, median, and range) inspected and compared with histological data in order to identify threshold values to define each biopsy-based gametogenic stage classification. For females, square root transformed ova areas collected via both histological and biopsy analyses were compared using the two sample student’s t-test and trends were identified between transformed ova areas and gametogenic classifications. Untransformed areas were evaluated in order to determine cell classification threshold values by approximating the midpoint value between the median oocyte areas of histological and biopsy-based distributions. Attribute agreement analysis (Minitab

Software v. 17.2.1) was conducted in order to compare gametogenic assessment data generated by the histological approaches, as well as the biopsy-based methods for both male and female samples. Gametogenic stage agreement between analytical methods was assessed using Cohen’s

Kappa Statistic and scores were interpreted based upon the Kappa statistic strength of agreement scale from Landis and Koch (1977) (Table 2.4).

Results

Male Histological Analyses

A total of 22 male E. complanata (n = 11 per site) were sampled and prepared for histological analysis of the gonad. No evidence of either intersex condition or functional hermaphroditism was observed in any histological sections examined. Total organism mass

(valves, mantle and visceral mass) ranged from 25.9 g to 91.2 g (x ̅ = 55.3 g; SD = 16.1 g) with 95%

CI = ± 7.2 g, while valve length ranged from 64.9 mm to 92.4 mm (x ̅ = 80.7 mm; SD = 6.5 mm) with

95% CI = ± 2.9 mm. These lengths, in light of studies on male unionid length/maturity relationships

(Haag and Stanton, 2003) are supportive of male sexual maturity for all sampled individuals.

Area-based histological analysis of male E. complanata sampled supported a seven-stage gametogenesis classification scheme, modified from Barber (1996) and Yokely (1972) (Table 2.5).

The first, an Inactive stage, is intended for samples from which few, if any, identifiable gametes can be detected and no evidence of recent resorption is observed. No samples were classified into this stage in the study. Second, a Spent/Resorbing (Image 2.1) stage is intended for samples in which viable spermatogenic cells may be observed, but hemocytes, phagocytes, and other atretic cells and cellular debris occupy, on average, ≥ 20 % of the acinar area measured. This value was selected as threshold for Spent/Resorbing categorization following initial qualitative visual inspection of samples and subsequent area-based histological analysis. The initial qualitative analysis identified only two individuals displaying Spent/Resorbing characteristics, as they each

16 appeared to exhibit a high degree of tissue resorption in the gonad. In order to quantitatively distinguish Spent/Resorbing individuals from other classifications, a midpoint value was calculated between, (1) the lowest average percentage of atretic tissue observed among the two qualitatively determined Spent/Resorbing organisms and (2) the maximum observed average percentage of atretic tissue measured among all other samples. This resulted in a value of 18.4 %.

A threshold for positive Spent/Resorbing classification was approximated by rounding the calculated midpoint to the nearest 5% multiple. This resulted in a threshold level of ≥ 20 % average atretic tissue for Spent/Resorbing categorization (Figure 2.1). Minimum and maximum average percent tissue data used to approximate threshold values utilized for gametogenic categorization are presented in Table 2.6.

Among the remaining organisms sampled, two separate and distinct spermatogenic pathways were evident, distinguished by the proportion of atypical tissue present in the gonad. A low rate of atypical spermatogenesis was detected in five of the ten organisms sampled in May

2013. In order to select a threshold value for future classification of samples, the midpoint value between the maximum observed average percentage of atypical tissue in May samples and the minimum observed average percentage of atypical tissue in August samples was calculated

(Figure 2.2) and a threshold for classification was approximated by rounding the calculated midpoint up to the next 5% multiple. This resulted in a threshold level of 25% (by area) atypical tissue in the gonad, at or above which results in classification into the atypical category of development. This atypical categorization was further divided into two stages, (1) Early Atypical

(Image 2.2), in which spermatozoa account for < 1 % of all acinar area measured, and (2) Late

Atypical (Image 2.3) in which spermatozoa account for ≥ 1 % of all acinar area measured. A very strong association was observed between spermatogenic pathway and sampling date (Fisher’s exact test: p = 0.0000060, α = 0.05), as 11 of 12 (one resorbing stage) organisms sampled in late

July/early August exhibited evidence of active atypical spermatogenesis, while nine of 10 organisms (one spent/resorbing stage) sampled in May were found to be actively developing via the typical spermatogenic pathway (Figure 2.2).

Typical development was assigned to samples exhibiting active gametogenesis with < 25

% average atypical tissue and < 20 % average atretic/resorbing tissue occupying acini within the gonad. This classification of development included three stages; (1) Early Active (Image 2.4), (2)

Late Active (Image 2.5), and (3) Mature (Image 2.6). The Early Active stage was characterized by the presence of immature cells (spermatocytes and spermatids) in the absence of mature spermatozoa. The Late Active stage was characterized by presence of both immature cells and mature cells, however spermatocytes and spermatids were observed at a higher combined estimated frequency than mature spermatozoa. Individuals were assigned to the Mature stage of typical development if mature spermatozoa accounted for a higher estimated cell frequency than spermatocytes and spermatids, combined.

The criteria and step-wise process for gametogenic classification by histological analysis is depicted in Figure 2.3. Based upon this classification scheme, six of ten (60 %) male E. complanata sampled in Spring 2013 were categorized as Late Active, while 30 % (three of ten) and

10% (one of ten) were categorized as Mature and Spent/Resorbing, respectively. For the Summer

2013 samples, this distribution had shifted to the greatest number of organisms exhibiting Late

Atypical development (50 %; total n = 12) while 42 % and 8 % were categorized as Early Atypical and Resorbing (Figure 2.4). Of those organisms sampled, no individuals met the criteria for

Inactive or Early Active classification.

Transect-Based Histological Approach Performance

Both the transect-based and tissue area-based approaches to histological analysis generated estimated cellular fractions for each gametogenic cell type and tissue included in the analysis. Sample reclassifications according to transect-based data compared to original area- based classifications are presented in Figure 2.5. Attribute agreement analysis results in a Cohen’s

Kappa Statistic of 0.70 between the area- and transect-based approaches to gametogenic staging in this sample, which suggests a “Substantial” degree of agreement between methods (Table 2.4).

Male Biopsy Method Performance

Viable gametogenic tissue was observed in 100 % of biopsies inspected. Proportions for each cell type (mature, developmental, atypical, atretic/other; each depicted in Image 2.7) were calculated and compared with data collected via the histological approaches described above.

Classification criteria were developed for the biopsy-based method to match the seven-stage classification scheme developed using the area-based histological approach (Table 2.7). As was the case for the area-based histological analysis, biopsy data were initially analyzed in order to determine a threshold value of atretic tissue to distinguish organisms undergoing active gametogenesis versus an atretic or resorptive condition (Figure 2.6). A threshold value of 90 % average atretic biopsy tissue, at or above which results in Spent/Resorbing classification, was selected by calculating the approximate midpoint value between the maximum and minimum observed average percentages of atretic biopsy tissue among samples grouped according to prior histological categorization as either Non-Spent/Resorbing or Spent/Resorbing, respectively. The same approach was taken for typical versus atypical tissue in the biopsy, resulting in a threshold value of 1.5 %, at or above which individuals were categorized as Atypical (Figure 2.7). The two stages within the atypical pathway category were differentiated by whether the largest fraction

19 of viable gametes observed in the sample was associated with mature spermatozoa (Late

Atypical) or some other gametogenic stage (Early Atypical). The remaining stage categories followed a classification pattern similar to the area-based histology approach and were based on presence/absence of gametogenic cells and observed frequencies of each.

Criteria for biopsy-based gametogenic staging are depicted in Figure 2.8. Sample reclassifications according to biopsy-based data were compared to (1) original area-based classifications (Figure2.9) and (2) transect-based classifications (Figure 2.10). Attribute agreement analyses result in Cohen’s Kappa Statistics of 0.88 between the area-based and biopsy-based approaches to gametogenic staging in this sample, and 0.68 between the transect-based and biopsy-based approaches to staging in this sample. These scores indicate “Substantial” staging agreement between transect and biopsy methods and “Almost Perfect” agreement between the biopsy method and the original area-based histological procedure (Table 2.4).

Female Histological Analyses

A total of 17 female E. complanata were sampled and prepared for histological analysis of the gonad. No evidence of either intersex condition or functional hermaphroditism was observed in any histological sections examined. Total organism mass (valves, mantle, and visceral mass) ranged from 28.1 g to 111.2 g ( x ̅ = 55.6 g; SD = 21.4 g) with 95% CI = ± 11.0 g, while valve length ranged from 69.0 mm to 91.9 mm (x ̅ = 79.6 mm; SD = 7.5 mm) with 95% CI = ± 3.8 mm, suggesting all organisms observed were of reproductive maturity (Downing et al., 1993).

Gametogenic stage was assigned to female E. complanata according to the classifications listed in Table 2.8. Criteria for each listed stage were based upon the proportions of developmental tissue, mature tissue, and atretic/resorbing tissue measured in each of 10 acini per organism (2 sections; n = 5 acini per section), as well as on previous descriptions (Barber 1996).

Based upon this classification scheme, six of ten (60 %) female E. complanata sampled in Spring

2013 were categorized as Mature (Image 2.8), while 30 % (three of 10) and 10 % (one of 10) were categorized as Developing (Image 2.9) and Spent/Resorbing (Image 2.10), respectively. For the

Summer 2013 samples, this distribution had shifted to Spent/Resorbing (100 %; total n = 7) as the only observed gametogenic stage (Figure 2.11). Of those organisms sampled, no individuals met the criteria for Inactive classification.

Untransformed ova area distributions collected via both the histology and biopsy procedures are shown in Figure 2.12. Untransformed histology-based ova areas for E. complanata classified histologically as Mature ranged from 633 µm2 to 8603 µm2, (median value = 3810 µm2), while biopsy-based ova areas for the same individuals encompassed a broader range, from 1088

µm2 to 25543 µm2 (median value = 7958 µm2). Significant differences were found between square root transformed ova areas of individuals classified as Mature and Developing using both the histological approach (2-sample student’s t-test; p = 0.034 ; α = 0.05) and the biopsy-based approach (2-sample student’s t-test; p < 0.0001 ; α = 0.05). Significant differences were also detected for square root transformed ova areas when the histological and biopsy-based data were compared, regardless of gametogenic stage classification (2-sample student’s t-test; p < 0.0001;

α = 0.05 for both classifications, Developing and Mature). Back transformed mean ova areas for the two methods and gametogenic stages are presented with 95% confidence intervals in Table

2.9. For individuals classified histologically as Spent/Resorbing, detection of viable ova in biopsy samples was rare and samples mainly appeared to contain non-viable oocytes, other atretic tissues, and cellular debris. Histological analysis of those ova areas observed in organisms classified Spent/Resorbing produced distributions much lower and more limited in range than those observed in Developing and Mature individuals, as the range of untransformed areas spanned from 294.1 µm2 to 2077.6 µm2, (median value = 1048.2 µm2) using histological methods

21 and from 25.3 µm2 to 3654.9 µm2, (median value = 1170.9 µm2) using the biopsy-based approach

(Figure 2.13).

Female Biopsy Method Performance

Viable gametogenic tissue was observed in approximately 88 % of female biopsies inspected (Image 2.11). Two samples contained only evidence of gonad resorption with no viable ova. For the remaining samples, ova areas were measured and compared with data collected via the histological approach described above. Classification criteria were developed for the biopsy- based method to match the four-stage classification scheme developed using the area-based histological approach (Table 2.8). Biopsy-based ova area data were initially analyzed in order to determine threshold area values to distinguish mature, developing, and atretic ova (Table 2.10).

These values were selected by approximating the midpoint between the median biopsy-based areas for organisms previously classified as Developing and Mature, as well as between those classified as Developing and Spent/Resorbing. Ova found to be below the midpoint of Developing and Spent/Resorbing groups (2974 µm2) were assigned to the Spent/Resorbing condition. In addition, ova observed to be clearly undergoing resorption and cellular breakdown, regardless of area measured, were also assigned to this category. Those ova found to be at or above the midpoint of Developing and Mature (6368 µm2) were classified as Mature while those below that value but at or above the Spent/Resorbing threshold were considered Developing. Ova frequencies for each classification observed in each biopsy sample were tabulated and a percentage for each cell stage was determined. The classification represented by the highest percentage in each sample dictated the biopsy-based gametogenic stage assigned.

Sample reclassifications according to biopsy-based data were compared to original histological area-based classifications (Figure 2.14). Attribute agreement analyses result in

Cohen’s Kappa Statistics of 0.80 between the area-based and biopsy-based approaches to gametogenic staging in this sample. This score represents the highest value to be classified as

“Substantial” staging agreement between the biopsy method and the original area-based histological procedure and falls just below “Almost Perfect” agreement between these methods

(Table 2.4).

Discussion

The nonlethal biopsy procedure for staging gametogenesis and assessing reproductive condition in freshwater mussels exhibited a high rate of agreement with traditional histological approaches for male E. complanata in this study and demonstrates the potential for application in other unionid species. Among the individuals sampled, a clear distinction was observed for male gamete development between the two sampling events. Male E. complanata collected in May

2013 exhibited generally synchronous development and appeared to be maturing prior to a late spring or early summer spawning period. For these individuals, full agreement in gametogenic staging was observed between the area-based histological approach and the alternative biopsy procedure, and all were found to be undergoing typical spermatogenesis. Within this spermatogenic pathway, spermatogonia develop within the acinar wall and, upon further development, differentiate into primary spermatocytes and are released into the acinar lumen. A sequential migration of cell stages is often evident in male acini as primary spermatocytes are found closest to the margins of the acinar lumen and subsequent stages form inward-moving bands or groups, from secondary spermatocytes to spermatids and, ultimately, mature spermatozoa. This spatial development appears to have an impact on the performance of the alternative, transect-based histological procedure utilized for comparison in this study, as is discussed later.

A number of different gametogenic classification schemes appear in the literature for bivalves that include separate “Spent” and “Resorbing” or “Atretic” stages of development

(Dinamani, 1974; Barber, 1996). Here, these stages were combined into a single Spent/Resorbing stage, as a primary characteristic often differentiating these two stages (collapse of the acinar wall), was unidentifiable in collected biopsies. As an atretic or resorptive stage often follows periods of spawning in order to clear acini of unreleased gametogenic tissues, the combination of these two potentially separate and distinct stages was justified.

Among males collected for analysis during the second sampling event in late July/early

August 2013, synchronous development was, again, generally observed, though the individuals sampled at the time appeared to be undergoing spermatogenesis through an alternative, atypical pathway. This “atypical” spermatogenesis has been observed among many unionid species and has been described in detail (Shepardson et al., 2012). Atypical spermatogenesis, to date, has been either indistinctly classified from typical spermatogenic tissue as Developing or Mature

(Beasley et al., 2000), or has been largely ignored in the development of gametogenic staging criteria (Cek and Sereflisan, 2011). As the full pathway of atypical development has been described and appears to result in the production of spermatozoa indistinguishable from those developed via the typical spermatogenic pathway, inclusion of this process in gametogenic staging criteria is warranted. Whether the mature gametes resulting from atypical gametogenesis serve an active reproductive role as is understood for typical spermatozoa is not yet known. Alternative hypotheses for the potential function of these cells have been proposed and range from providing nutrition to the egg to acting in the formation of spermatozeugmata (large, membrane encapsulated aggregates of mature spermatozoa formed prior to spawn in many mussel species)

(Shepardson et al., 2012). In this study, the appearance and apparent maturation of these cells

24 seemed to generally align with gametogenic resorption in females sampled concurrently, and so, herein, suggests a possible alternative role to typical male reproductive function.

While all male cell types were observed in this study, the earliest of the spermatogenic cells, spermatogonia, were not included in the development of staging criteria. Collection of spermatogonia during biopsy sampling appeared low relative to expected frequencies. It is hypothesized that this is a function of their physical relationship with the acinar wall and the nature of the biopsy procedure. As a small amount of distilled water is injected into the gonad and the resulting gonad fluid is retracted, unbound cells located within the acinar lumen appear to have a greater chance of collection than those remaining physically attached to the acinar wall.

The small volume of water injected into the gonadal tissue may not, on average, exert a disruptive force great enough to dislodge attached and/or embedded cells. This hypothesis was supported by results of female sampling, as similar issues with a lack of immature cells, characterized by a close association with, and physical attachment to, the acinar wall, were observed among biopsies collected. During female unionid development, it is only during late maturation periods that the stalk-like formations attaching developing ova to the acinar wall release, allowing free oocytes to migrate into the acinar lumen (Cek and Sereflisan, 2011). A low frequency of identifiable developmental ova stages (oogonia, primary oocytes, etc.) in biopsies drawn from female E. complanata in this study suggests a potential limitation of the biopsy procedure in collecting fully representative samples of the cellular makeup of the gonad. Acknowledgement of this limitation, however, does not appear to influence its ability to stage gametogenesis significantly, although its effect on stages that were not observed in this study can only be speculated.

Though methodological limits are apparent, the biopsy-based approach to staging gametogenesis in unionids offers a number of advantages over traditional histological methods.

As was presented in Figures 2.3 and 2.8, the methods described here attempt to quantify reproductive tissues to guide staging decisions. While previous histological assessments have utilized quantitative approaches, such as transect-based cell counts, in studies of gamete development in various bivalves (Jones et al,. 1986; Haggerty et al., 1995; Garner et al., 1999), these methods have typically resulted in, or been aligned with, qualitative criteria for gametogenic stage assessments that can be open to interpretation between different investigators. The biopsy-based methodology described here provides a quantitative approach to staging that limits the potential for inter-observer differences to occur by defining quantified thresholds to categorize gametogenic stages. The quantitative analytical approach also made possible a direct comparison of two alternative histological procedures (area-based measures and transect-based cell counts) that have appeared in the literature to guide gametogenic stage assessment. In order to determine whether the simpler, transect-based approach to histological examination of the gonad in E. complanata was a viable alternative methodology for the initial determination of gametogenic stage, results generated using this approach were used to reclassify samples according to the area-based gametogenic classification scheme described earlier. The results of this comparison, as well as the benefits and limitations of each histological approach, were used to guide selection of a reference method for biopsy comparison. As attribute agreement analysis suggested “Substantial” agreement between the two histological approaches (Cohen’s Kappa = 0.70 out of a possible 1.0), several other factors were considered in selecting a reference method. The transect-based procedure is simple and requires less time to employ than the area-based method. In addition, because of the size differences that exist among the different spermatogenic cell stages, a relatively low number of (larger) cells in early stages of development compared with a relatively high number of (smaller) mature cells might be more accurately estimated using cell counts. However, as was described earlier, the nature of the

26 unique spatial organization within acini of freshwater mussels might influence the estimated relative abundance of the various cell stages present when a single transect stretching the greatest length and passing through the approximate center of the acinus is employed. In comparison, the utilization of an area-based procedure better represents the relative masses of tissue at each stage present within acini. However, again due to the relative size differences that exist between these cell stages, equal areas for a developmental stage and a mature stage do not equate to equivalent abundance of each cell type. This issue can be circumvented, however, when average area occupied by a single cell of each stage is determined, and area occupied is converted to an estimated number of cells present. These estimates, in addition to area measures for atretic and atypical tissues, resulted in consistent agreement with cell count data collected via the biopsy procedure when used to stage gametogenesis (Cohen’s Kappa score = 0.88). For these reasons, the area-based procedure was selected as the reference method for biopsy comparison in this study and results support the utility of this nonlethal procedure as an alternative to histological analysis in staging gametogenesis and assessing reproductive condition in male E. complanata throughout the reproductive cycle.

In comparison to that for males, the performance of the biopsy method was weaker for female E. complanata in this study. Traditional staging of gametogenesis in females was conducted following histological analysis, and reclassifications were performed based on data collected from biopsy sampling. Ova diameters (µm) have been typically utilized in studies of reproductive development and gametogenesis in bivalves (Barber 1996; Barber and Blake 1981;

Haggerty et al., 1995), however ova areas (µm2) were used for all analyses in this study. Barber

(1996) suggested the use of ova area measures in place of diameter in order to reduce variability of the measure, as non-spherical objects are often observed during histological analysis of the female gonad. The use of diameter measures on biopsy samples, in which complete,

27 unobstructed, spherical ova are typically observed result in acceptably accurate estimates of area

(data not shown), however due to the non-spherical and partially obstructed nature of most ova observed in histological samples of female gonad, the utility of area-based measures is suggested.

Using the staining procedures and microscopic analysis described earlier, distinguishing mature oocytes collected in the biopsy from developing cells that were dislodged from the acinar wall was not possible. However, based upon histological staging, ova areas measured in biopsy samples were classified as mature, developing, and atretic based upon threshold values obtained by comparing data from organisms histologically classified as Mature, Developing, and

Spent/Resorbing. No Inactive individuals were observed in this study, and so the ability of the biopsy method to classify such individuals is unknown.

The procedure described for a nonlethal evaluation of gametogenesis in the freshwater mussel, E. complanata shows strong agreement with information obtained through traditional histological analysis. The ability to gain such information about a unionid population could be of value in future conservation efforts, particularly for species considered threatened or endangered, as this provides the potential to gain information about gamete development with little to no organism sacrifice. Whether the specific methodology described in this study for E. complanata in the Cacapon River, WV is applicable to other populations and/or unionid species, however, is unknown.

Developmental Stage Spermatogenic Pathway Cell Descriptions

Largest of the male gametes (diameter approx. 8 um), roughly Spermatogonia spherical in shape and growing out of the follicle wall.

Spherical in shape and smaller than Spermatogonia. Diameters range Spermatocytes approximately 5 um (secondary) to 7 um (primary).

Shape variable, ranging from spherical (diameter approx. 4 um) to Spermatids ovular (maximum length approx. 4 to 5 um).

Cylindrical to bullet-shaped with visible flagellum. Generally lightly Spermatozoa basophilic. Maximum lengths ranged approx. 3.0 to 3.5 um.

Clustered masses of deeply basophilic spermatocytes (approx. 1 to 32

cells). May be found within sertoli cells protruding from the acinar wall Sperm Morulae or free in the acinar lumen. Terminal division of each cell within in the

mass results in the production of two mature spermatozoa.

Table 2.1 Spermatogenic cell classifications in male E. complanata modified from Cek and

Sereflisan, 2011 and McElwain and Bullard, 2014.

Developmental Stage Oogenic Pathway Cell Descriptions

Viable ova embedded within, or attached to, the acinar wall. Range

Immature Oocytes from small, fully embedded primary oogonia to post-vitellogenic

oocytes with intact stalk-like attachment point.

Vitellogenic oocytes surrounded by loose, outer vitelline membrane

Mature Oocytes that have detached from the acinar wall and migrated toward the

interior of the acinus.

Table 2.2 Oocyte cell classifications in female E. complanata modified from Cek and Sereflisan,

2011.

Developmental Stage Spermatogenic Pathway Cell Descriptions

Largest of the typical male gametes (diameter approx. 8 um), roughly Spermatogonia spherical in shape.

Spherical in shape and smaller than Spermatogonia. Diameters range Spermatocytes approx. 5 to 7 µm.

Shape variable, ranging from spherical (diameter approx. 4 µm) to Spermatids ovular (maximum length of approx. 4 to 5 µm).

Cylindrical to bullet-shaped with visible flagellum. Generally lightly Spermatozoa basophilic. Maximum lengths ranged approx. 3.0 to 3.5 µm.

Highly basophilic, multicellular bodies ranging in diameter from Sperm Morulae approx. 5 to 15 µm.

Table 2.3 Biopsy-generated spermatogenic cell classifications in male E. complanata

Kappa Statistic Strength of Agreement

< 0.00 Poor

0.00 – 0.20 Slight

0.21 – 0.40 Fair

0.41 – 0.60 Moderate

0.61 – 0.80 Substantial

0.81 – 1.00 Almost Perfect

Table 2.4 Cohen’s Kappa Statistic Strength of Agreement Scale

Gametogenic Stage Histology-based Male Stage Descriptions

No identifiable gametes observed and follicles non-existent or elongate. (0) Inactive Walls characterized by undifferentiated germinal epithelium. Immature developmental cells (spermatocytes, spermatids) constitute (1) Early Active the greatest estimated frequency; no mature spermatozoa present. Sperm morulae occupy < 25 % of total acinar area measured. Immature cells combined (spermatocytes and spermatids) constitute (2) Late Active the greatest estimated frequency; spermatozoa present. Sperm morulae occupy < 25 % of total acinar area measured. All cell stages generally present; mature spermatozoa comprise the (3) Mature greatest estimated frequency of cells observed. Sperm morulae occupy < 25 % of total acinar area measured. Hemocytes, phagocytes, and other atretic cells and cellular debris (4) Spent/Resorbing occupy ≥ 20 % of acinar area measured; viable spermatogenic cells may also be present at any combination of levels. Sperm morulae occupy ≥ 25 % of all acinar areas measured. (5) Early Atypical Spermatocytes typically present; spermatozoa generally absent, and occupy < 1% of total acinar area. Typical spermatids generally absent. Sperm morulae occupy ≥ 25 % and spermatozoa occupy ≥ 1 % of total (6) Late Atypical acinar area measured. Spermatocytes may be present.

Table 2.5 Gametogenesis classifications in male E. complanata, modified from Barber (1996) and

Yokely (1972).

Image 2.1 – Acinus (40x objective) representing Spent/Resorbing gametogenic classification in male E. complanata. Acinus has shrunken appearance and actively resorbing cells, and infiltration of hemocytes (HC) is evident. Gametes (developing or mature) appearing viable may also be observed, though most are undergoing active resorption via phagocyte (PC) activity.

t 20

Non-Atretic Atretic Initial Qualitative Classifications

Figure 2.1 Average percentage values for proportion of atretic tissue observed in histological sections of gonad tissue of male E. complanata collected in May and late July/early August 2013.

Dashed line indicates threshold level (20%) selected to define Non-Atretic (areas below line) versus Atretic (areas at or above line) classification and categorization as Spent/Resorbing.

100

n 80

s 60

p 40

g 20

Spring '13 Summer '13 Sample Date

Figure 2.2 Average percentage values for proportion of atypical tissue observed in histological sections of gonad tissue of male E. complanata collected in May and late July/early August 2013.

Dashed line indicates threshold level (25%) selected to define typical (areas below line) versus atypical (areas at or above line) categorization.

Image 2.2 – Acinus (40x objective) representing Early Atypical gametogenic classification in male

E. complanata. Large finger-like sertoli cells can be seen growing from the acinar wall and orienting toward the inner lumen. Both within these large cells and free in the acinar lumen can be seen sperm morulae (SM), membrane-bound spermatid aggregations of two to app. 32 cells, ranging in size and number of cells. Typical cells (spermatocytes, spermatids, spermatozoa) may be observed, though spermatozoa are found to occupy less than or equal to 1% of acinar area and atypical cells (morulae, sertoli cells) occupy ≥25% of the total acinar area.

Image 2.3 – Acinus (40x objective) representing Late Atypical gametogenic classification in male

E. complanata. Large finger-like sertoli cells can still be seen growing from the acinar wall and orienting inward toward the inner lumen. Sperm morulae (SM) are still common, and atypical cells still occupy ≥25 % of the total acinar tissue. Mature spermatozoa (SZ), in this case, occupy ≥

1% of the total area of the acinus.

Image 2.4 – Acinus (40x objective) representing Early Active gametogenic classification in male

E. complanata. Few (<20% by area), if any, atypical spermatogenic cells are present. No mature spermatozoa are present. Acinus is filled with spermatocytes (SC) and/or spermatids (SP).

Image 2.5 - Acinus (40x objective) representing Late Active gametogenic classification in male E. complanata. Few (<20% by area), if any, atypical spermatogenic cells, such as sperm morulae

(SM) are present. Mature spermatozoa are present, but are estimated to be less numerous than the estimated number of spermatocytes (SC) and spermatids (SP) combined.

Image 2.6 - Acinus (40x objective) representing Mature gametogenic classification in male E. complanata. Few (<20% by area), if any, atypical spermatogenic cells, such as sperm morulae are present. Mature spermatozoa are present, and are estimated to be more numerous than the estimated number of spermatocytes (SC) and spermatids (SP) combined.

Minimum Maximum Calculated Measured Cell/ Selected Threshold Category/Stage % Among % Among Min/Max Tissue Type for Classification Classified Unclassified Midpoint

Spent, Atretic/Resorbing 25.4 % 11.4 % 18.4 % ≥ 20 % Resorbing Tissues

Atypical Pathway Atypical 45.2 % 0.7 % 22.3 % ≥ 25 % Tissues

Table 2.6 – Threshold value approximations for male E. complanata used in the determination of area-based histological analysis.

Evidence of Active or Resorbing Gonadal Tissue

No Yes

Classify Average % Inactive Atretic Tissue

≥ 20 % < 20 %

Classify Average % Spent/Resorbing “Atypical” Tissue

≥ 25 % < 25 %

Average % Estimate # Spermatocytes (SC), Spermatozoa Spermatids (SP), Spermatozoa (SZ)

< 1 % ≥ 1 % SZ = 0 SZ < SC + SP

Classify Classify Classify Classify Early Atypical Late Atypical Early Active Late Active

SZ ≥ SC + SP

Classify Mature

Figure 2.3 – Histology-based male freshwater mussel gametogenic classification protocol generated utilizing E. complanata from the Cacapon River, WV.

Spring '13 Summer '13 60

)

(

e 40

e 20

O 10

e e g l l e e g l l r a a r a a iv n c c iv n c c t u i i i t u i i i t b t b c a p p c a p p r y y r y y A o t t A o t t M s M s e A A e A A t e t e a R y e a R y e L l t L l t r a r a a L a L E E

Panel variable: Sample Date Area-Based Gametogenic Stage Percent is calculated within all data.

Figure 2.4 – Gametogenic stage observed for male E. complanata in Spring (May) 2013 and

Summer (late July/early August) 2013 based upon area-based histological analysis. Figure frequencies represent analysis of 10 (Spring ’13) and 11 (Summer ’13) male specimen.

Area-Based Gametogenic Stage Transect-Based Gametogenic Stage 90 80

v 70

s 60

O 50

n 40

r 30

P 20 10 0 Stage e e l l e e l l e e l l e e l l v r g a a v r g a a v r g a a v r g a a i u in ic ic i u in ic ic i u in ic ic i u in ic ic ct t b p p ct t b p p ct t b p p ct t b p p A a r y y A a r y y A a r y y A a r y y M so t t M so t t M so t t M so t t te e A A te e A A te e A A te e A A a R l y e a R ly e a R ly e a R ly e L t/ r at L t/ r at L t/ r at L t/ r at n a L n a L n a L n a L e E e E e E e E Sp Sp Sp Sp Date 13 13 13 13 ' ' ' ' g r g r n e n e ri m ri m Sp m Sp m Su Su

Panel variable: Staging Method Percent is calculated within levels of Date.

Figure 2.5 - Male E. complanata gametogenic stage classifications as generated by the area- based and transect-based histological approaches (total n Spring + Summer = 21)

Image 2.7 – Biopsy (40x objective) of male gonad from E. complanata showing mature spermatozoa

(SZ), spermatocytes (SC), spermatids (SP) and an atypical pathway morulae (SM). Non-spermatogenic cells and cellular debris contributing to the “atretic/other” category are also seen in this biopsy.

100

p 60

c 50

t 40

% 30

Non-Atretic Atretic Histological (Area-Based) Gametogenic Classification

Figure 2.6 – Individual value plot indicating threshold value for atretic (≥ 90%) and non-atretic

(< 90%) categorization in male biopsy analysis (n = 21 organisms sampled)

y 8

o 7

l 6

p 5

% 3

Typical Atypical Histologcal (Area-Based) Gametogenic Classification

Figure 2.7 - Individual value plot indicating threshold value for Typical and Atypical categorization in male biopsy analysis (n = 21).

Evidence of Active or Resorbing Gonadal Tissue

No Yes

Classify Average % Inactive Atretic Tissue

≥ 90 % < 90 %

Classify Average % Spent/Resorbing “Atypical” Tissue

≥ 1.5 % < 1.5 %

Average % Spermatocytes (SC), Average % Spermatocytes (SC), Spermatids (SP), Spermatozoa (SZ) Spermatids (SP), Spermatozoa (SZ)

SZ ≥ SC + SP SZ < SC + SP SZ < SC + SP SZ = 0

Classify Classify Classify Classify Late Atypical Early Atypical Late Active Early Active

SZ ≥ SC + SP

Classify Mature

Figure 2.8 - Biopsy-based male freshwater mussel gametogenic classification protocol generated utilizing E. complanata from the Cacapon River, WV

Area-Based Gametogenic Stage Biopsy-Based Gametogenic Stage 70

d 60

v r 50

s b 40

n 30

e 20

P 10

0 Stage e e l l e e l l e e l l e e l l v r g a a v r g a a v r g a a v r g a a i u in ic ic i u in ic ic i u in ic ic i u in ic ic ct t b p p ct t b p p ct t b p p ct t b p p A a r y y A a r y y A a r y y A a r y y M so t t M so t t M so t t M so t t te e A A te e A A te e A A te e A A a R l y e a R ly e a R ly e a R ly e L t/ r at L t/ r at L t/ r at L t/ r at n a L n a L n a L n a L e E e E e E e E Sp Sp Sp Sp Date 13 13 13 13 ' ' ' ' g r g r n e n e ri m ri m Sp m Sp m Su Su

Panel variable: Staging Method Percent is calculated within levels of Date.

Figure 2.9 - Male E. complanata gametogenic stage classifications as generated by the area- based histological method and biopsy-based analysis (total n Spring + Summer = 21).

Biopsy-Based Gametogenic Stage Transect-Based Gametogenic Stage 90 80

v 70

s 60

O 50

n 40

r 30

Panel variable: Staging Methods Percent is calculated within levels of Date.

Figure 2.10 - Male E. complanata gametogenic stage classifications as generated by the transect-based histological method and biopsy-based analysis (total n Spring + Summer = 21).

Gametogenic Stage Biopsy-based Male Stage Descriptions

Few, if any, identifiable gametes observed (<25 cells/stage in biopsy) (0) Inactive and no evidence of actively resorbing gametogenic tissues.

Immature cells combined (spermatocytes, spermatids) constitute the

(1) Early Active largest cell fraction; no mature spermatozoa present. Sperm morulae

comprise < 1.5 % of all cells observed.

Immature cells combined (spermatocytes and spermatids) constitute

(2) Late Active the largest cell fraction; spermatozoa present. Sperm morulae

comprise < 1.5 % of all cells observed.

All cell stages generally present; mature spermatozoa comprise the

(3) Mature largest fraction of viable gametic cells observed. Sperm morulae

comprise < 1.5 % of all cells observed.

Hemocytes, phagocytes, and other resorbing cells and cellular debris

(4) Spent/Resorbing comprise more than 90% of cells observed; viable spermatogenic cells

may be present at any corresponding level(s).

Sperm morulae comprise ≥ 1.5 % of all cells observed. Spermatocytes

(5) Early Atypical typically present; spermatozoa may be present, but at a lower fraction

than other spermatogenic stages. Typical spermatids generally absent.

Sperm morulae comprise ≥ 1.5 % of all cells observed. Spermatocytes

(6) Late Atypical and sperm morulae present; spermatozoa comprise the largest fraction

of gametes observed.

Table 2.7 Biopsy-Generated Gametogenesis Classifications in male E. complanata

Gametogenic Stage Histology-based Female Stage Descriptions

Few, if any, identifiable gametes observed (<25 total in biopsy) and no (0) Inactive evidence of actively resorbing gametogenic tissues.

Immature cells (oogonia, developmental oocytes) occupy the largest

(1) Developing proportion of the gonad; mature (free) oocytes may be present at a

lower proportion.

Mature (free) oocytes occupy the largest average proportion of area (2) Mature within the gonad.

Hemocytes, phagocytes, and other resorbing cells and cellular debris

comprise a proportion of the acini greater than that of the developing

(3) Spent/Resorbing and mature cells combined. These cells may be present at any

corresponding level(s) that, combined, equal less area occupied than

atretic tissue.

Table 2.8 – Histology-based gametogenic stage classification for female E. complanata.

Image 2.8 – Acinus (40x objective) representing Mature gametogenic classification in female E. complanata. Mature ova occupy the greatest proportion of the acinus. Ova classified as

Developing and/or Atretic may or may not also be present. Mature ova have detached from the acinar wall and pack into the acinar lumen, prior to their transfer to the water tubes for potential fertilization.

Image 2.9 – Acinus (40x objective) representing Developing gametogenic classification in female

E. complanata. Immature ova (here identified by size and apparent attachment to the acinar wall) occupy the greatest proportion of the acinus. Ova classified as Mature and/or Atretic may or may not also be present. Mature ova have detached from the acinar wall and pack into the acinar lumen, prior to their transfer to the water tubes for potential fertilization, while immature ova retain a physical attachment to the acinar wall via a stalk-like structure.

Image 2.10 – Acinus (40 x objective) representing Atretic gametogenic classification in female E. complanata. Atretic or resorbing ova and other cells of resorption occupy a greater proportion of the acinus than developing and mature ova, combined. Ova classified as Mature and/or

Atretic may or may not also be present. The acinus is generally shrunken, and most gametogenic tissue unreleased during the previous spawning period appears to be undergoing resorption.

Spring '13 Summer '13 100

b 60

r 40

0 Developing Mature Spent/Resorbing Developing Mature Spent/Resorbing

Female Stages Panel variable: Date 2 Percent is calculated within all data.

Figure 2.11 – Gametogenic stages observed for female E. complanata in Spring (May) 2013 and

Summer (late July/early August) 2013 based upon histological analysis.

Histology Biopsy

Mean (µm2) 95% CI (µm2) Mean (µm2) 95% CI (µm2)

Developing Ova 2695.7 3069.2 – 2342.6 5353.8 6448.1 – 4369.2

Mature Ova 3571.3 3831.6 – 3317.8 8385.1 9082.1 – 7726.4

Table 2.9 – Back-transformed mean and 95% Confidence Values determined following square root transformations of ova measured in histological and biopsy samples from female E. complanata according to gametogenic stage assigned.

Image 2.11 – Biopsy of female gonad (4x objective) from E. complanata showing mature (M), developing (D) and atretic (A) ova. Designations were based on threshold areas generated by histological analysis as well as staining characteristics and appearance.

Developing Mature Method Histology 0.00030 Biopsy

0.00025

0.00020

n e 0.00015

0.00010

0.00005

0.00000 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 4 8 2 6 0 4 4 8 2 6 0 4 1 1 2 2 1 1 2 2 Measured Ova Areas (µm2)

Panel variable: Stage 2_1

Figure 2.12 – Ova area distributions (untransformed) for developing and mature ova observed in

E. complanata samples according to measurement method.

Histology Areas (um) Resorbing Biopsy Areas (um) Resorbing 50

b 30

c r 20

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 5 0 5 0 5 0 5 5 0 5 0 5 0 5 1 1 2 2 3 3 1 1 2 2 3 3 Ova Area (µm2)

Figure 2.13 – Untransformed ova area distributions resulting from histological and biopsy-based analysis for E. complanata classified as Spent/Resorbing

Gametogenic Stage Biopsy Ova Area Threshold (µm2)

Spent/Resorbing < 2974

Developing ≥ 2974, <6368

Mature ≥ 6368

Table 2.10 – Threshold values determined for biopsy-based identification of gametogenic stage in female E. complanata based on ova areas observed in collected biopsy fluid.

Histology Biopsy 100

e 80

b 60

c 40

P 20

0 Stage g re g g re g g re g g re g in u in in u in in u in in u in p t b p t b p t b p t b o a r o a r o a r o a r l M o l M o l M o l M o ve es ve es ve es ve es e R e R e R e R D D D D

Season g r g r n e n e ri m ri m p m p m S u S u S S

Panel variable: Histo Biopsy Appraiser Percent is calculated within levels of Season.

Figure 2.14 – Female E. complanata gametogenic stage classifications as generated by histological and biopsy-based analysis.

Chapter 3: Laboratory and field investigations into estrogenic effects on growth and

gametogenesis in freshwater mussels

Introduction

In recent years, a general decline in aquatic biodiversity has become evident, as exemplified by the current plight of freshwater mussels, one of North America’s most endangered faunal groups (Augspurger et al., 2007; Haag and Williams, 2014). Proposed factors contributing to this decreased diversity of freshwater mussels, as well as fish, crayfish, and amphibians, include such factors as stream fragmentation, flow regulation, channelization, dredging, sediment loading, and pollution from land-use activities and discharge of toxic contaminants from agricultural, municipal, and industrial sources (Ricciardi, 1999, Haag & Williams, 2014). Significant effects of xenobiotic exposure on reproduction and development in freshwater invertebrates have been demonstrated in the literature (Hardege et al., 1997; Jobling et al., 2004; Nentwig,

2007; Ortiz-Zarragoitia and Cajaraville, 2006), and the effects of a particular group of compounds, termed endocrine disrupting chemicals (EDCs), have been implicated in this trend (Gagné et al.,

2001; Gagné and Blaise, 2003; Regoli et al., 2001). Studies have identified a number of such compounds to date, both natural and synthetic, which can mimic the effects of steroidal hormones (androgens and estrogens) that are critical to reproduction and development of many organisms (Farris and Van Hassel, 2007; Landis and Yu, 2004). The sources of these compounds vary widely, from municipal wastewater treatment and industrial effluents to agricultural practices of pesticide application and livestock waste utilization. However, information on the impacts of this group of compounds on wildlife populations is limited to a relatively small number

64 of species, and knowledge of the responses and relative sensitivities of different animal species to such compounds is lacking (Jobling et al., 2004). While it is important to understand the underlying mechanisms of activity associated with compounds such as EDCs, evaluating adverse effects of such chemicals on the development and reproduction of potentially affected populations is also an important goal and should not be delayed (Gourmelon and Ahtiainen,

2007).

A variety of land-use practices are known to have the potential to contribute non-point source pollution to surface waters, including estrogens and other EDCs. Therefore, investigations into the health and conservation status of natural unionid populations should consider land-use practices in the assessment and identification of factors that might influence the ultimate success or failure of these organisms. The approximately 1761 square kilometer area comprising the

Cacapon River Basin in West Virginia (WV) is located within what is considered a central location of poultry production (Chambers and Leiker, 2006). The drainage area of the Cacapon’s primary upper tributary, the Lost River, has the highest poultry house density in the country, which raises significant water quality concerns as poultry litter waste is typically land-applied (Elrashidi et al.,

2008). Several estrogenic, reproductive hormones and their metabolites are excreted into poultry litter, including, for example, 17β-estradiol (Nichols et al., 1997; Finlay-Moore et al., 2000). This hormone, and its primary metabolite estrone (E1) are naturally occurring vertebrate hormones that are excreted by chickens, primarily in the urine, in the range of 2.5 µg – 6 µg per individual female per day (Combalbert and Hernandez-Raquet, 2010). As more than 90,000,000 birds were reared and sold in the region of the Cacapon River in 2002 (Chambers and Leiker, 2006), and the majority of the litter accumulating this estrogen load is locally applied, this industry serves as a significant potential source of estrogens in the local environment. Although lab biodegradation and transformation studies have shown that estrogens are biodegraded by many different types

65 of organisms, and column experiments have indicated that estrogens may not easily leach through soil, field studies have demonstrated that manure based estrogens are sufficiently mobile to impact both surface and groundwater quality (Hanselman et al., 2003). In addition to the non- point contamination of ground and surface waters that may occur as a result of agricultural practices in the Cacapon River drainage, at least 24 NPDES (National Pollution Discharge

Elimination System) discharge permits have been granted by the West Virginia Department of

Environmental Protection in the area, and a large, though unquantified portion of residential sewage in the Cacapon River Basin may be inadequately treated or discharged directly to surface waters (Chambers and Leiker, 2006). This gives rise to the potential for environmental contamination, particularly by way of municipal sewage effluent and runoff from pasture and crop field application of composted poultry house litter. The existence of such point and non-point sources of both ground and surface water contamination may result in intermittent and possibly chronic exposure of freshwater mussels to both natural (such as 17β-estradiol) and synthetic estrogenic EDCs, as this river supports a number of different freshwater mussel species, the most common being the eastern elliptio, E. complanata (Villella and Smith, 2005). This type of exposure has been implicated in the disruption of sexual differentiation processes and inhibition of normal development of the gonad in E. complanata (Gagné et al., 2001).

Information on unionid and other molluscan responses to estrogen, xenoestrogen, and other organic contaminant exposure is limited, however the body of ecotoxicological research on these organisms has grown in recent years. Adults of the freshwater mussel Lampsilis fasciola exhibited significant behavioral changes in both males and females, as well as reduced energy reserves and altered metabolite production following 12-day exposures to either 17α- ethinylestradiol (EE2) or the aromatase inhibitor, fadrozole hydrochloride, at environmentally relevant concentrations (Leonard et al., 2014a; Loenard et al., 2014b). Following tricyclic

66 antidepressant (TCA) exposure, Hardege et al. (1997) observed inhibitory effects on spawning, fertilization, and development of embryos in the zebra mussel, Dreissena polymorpha, while Peck et al. (2007) concluded that this species could be susceptible to estrogenic contaminant exposure following analysis of the uptake, metabolism, and bioconcentration of E2 in tissues. Municipal and industrial effluent exposures have also provided evidence for a link between estrogenic compounds in the environment and endocrine or reproductive effects in mussels (Gagné et al.,

2001; Jobling et al., 2004). Keller et al. (2007) provide an excellent review of sublethal effects observed in unionids following exposure to various pesticides and other organics, and, though it is conceded that significantly less work has been undertaken on unionid responses to these compounds than metals, they conclude that the presence of these compounds in the environment poses a significant risk to mollusc populations.

The waters comprising the Cacapon watershed ultimately flow into the Chesapeake Bay, the Nation’s largest estuary, which has been impacted by degraded water quality, loss of habitat, and declines in biological communities (Philips, 2005), and recent evidence suggests that degraded water quality may be impacting the aquatic biota of the Cacapon River system, as well

(Chambers and Leiker, 2006). A study conducted by the United States Geological Survey (USGS) investigated the prevalence of intersex condition (i.e., development of eggs in the male gonad) in smallmouth bass populations throughout the Potomac Drainage, following observations of the condition during a 2003 investigation of fish kills in the area (Blazer et al., 2007). In a separate study conducted by USGS, passive sampling devices (PSDs) were deployed throughout the

Potomac Drainage in order to determine the presence of emerging contaminants, including several known and suspected EDCs (Chambers and Leiker, 2006). As a result of that study, several potential endocrine disrupting compounds were detected in samples taken from waters of the

Potomac Drainage, including the Cacapon River. However, natural and synthetic estrogens

67 typically associated with the agricultural land use activities and municipal effluents were not evaluated.

This work focuses on the potential influence of estrogens on gametogenesis and reproduction in unionids by assessing the extent to which estrogenic, potentially endocrine disrupting compounds occur seasonally in surface waters of the Cacapon River, WV while considering reproductive characteristics of resident E. complanata. The study seeks to test the following hypotheses:

H1: If the agricultural practice of poultry litter land application in the Cacapon River

watershed results in surface water contamination by the natural estrogen, 17β-

estradiol and its metabolites, effects of exposure on the resident freshwater

mussel, E. complanata will include the presence of intersex condition and/or other

reproductive impairment, as has been detected in smallmouth bass throughout this

drainage basin. In addition, the presence of these estrogens should result in

detectable levels of estrogenic activity of passive water samplers deployed in these

surface waters.

H2: If the primary vertebrate estrogen, 17β-estradiol (E2), plays an active role in unionid

gamete development and reproduction, then exposure of the freshwater mussel,

E. insulsa to exogenous E2 will result in a significant reproductive and/or

gametogenic response.

Four locations in the Cacapon River were preliminarily sampled; of these, two distinct populations of E. complanata were selected for further study, in areas primarily characterized by either agricultural land-use or forest. Reproductive characteristics associated with these organisms – specifically, gametogenesis, reproductive condition, prevalence of intersex, and male to female

68 ratios – were evaluated for individuals sampled from each population and characteristics of each population were compared. Additionally, a second unionid species, E. insulsa, was exposed in the laboratory to varying concentrations of the vertebrate estrogen, 17β-estradiol, and potential effects on gametogenesis and reproductive condition were explored.

Materials and Methods

Field studies with Elliptio complanata

Site Selection and Field Sampling

Four sites along the Cacapon River, West Virginia (WV), were located in August, 2008 for initial organism sampling ranging from the most upstream reaches to near the mouth and based upon accessibility and observed E. complanata abundance. Organism sampling was conducted via hang-grab and visual identification using glass-bottom buckets. E. complanata (n=25) from each site were collected, massed, and immediately sacrificed at the site in late September, 2008.

All excised tissues were fixed in 10% Neutral Buffered Formalin (NBF) for future histological processing. In the laboratory, a frontal section (approx. 0.5 mm) was cut from each visceral mass at the approximate midpoint of the anterior/posterior length. Samples were dehydrated in an ethanol (EtOH) series and paraffin embedded prior to sectioning and staging, and were stained with standard hematoxylin and eosin. Samples were analyzed to determine sex and to evaluate incidence of intersex and reproductive condition among organisms sampled at each site. Two of the four sites were selected for additional study. Site 1 (Images 3.1 and 3.2) was located at approximately 39° 05’ 31.1” N, 78° 35’ 42.9” W (elevation 291.4 m) in Wardensville, Hardy County,

WV, while Site 2 (Images 3.3 and 3.4) was located at approximately 39°36’26.31” N, 78°17’29.21”

(elevation 133.2 m), just downstream of a low-water bridge on Power House Road (CR 9/12) near

Berkeley Springs, Morgan County, WV. Sampling events occurred between September, 2009, and

October, 2013. During each sampling event, individuals were collected via hand-grab with aid of a glass-bottom bucket from each of the two Cacapon River sites. Organisms at Site 1 were found immediately off of the left descending bank in a shallow run exhibiting a high density of E. complanata. Organisms were observed more sporadically at Site 2, though most were collected from a mid-stream run. All collected individuals were massed and a subsample was subsequently biopsied for sex determination and/or gametogenic stage assessment. Additional organisms were sacrificed for histological analysis during May and July/August 2013 sampling events. In order to preserve tissues for analysis, both the biopsied tissue and the visceral mass of each sacrificed individual were immediately fixed in 10% NBF at the site. Gametogenic stage was determined for individuals biopsied during 2013 sampling events according to procedures described in Chapter 2

(see also Figure 2.8, Table 2.10). Histological analysis of organisms sampled in May and

July/August 2013 (n = 10/site) permitted calculation of two reproductive metrics, here termed the Acinar Area Fraction (AAF) and the Gonad Area Fraction (GAF). The GAF was modified from

Henley (2002) and, briefly, was calculated by taking the average proportion of area within the gonad containing developing or ripe tissues to the total gonad area observed throughout each of three fields of view (total magnification 4x). This metric provided a measure of the degree to which developing or mature acini filled the visceral mass. Similarly, the AAF was calculated as the proportion of area filled with viable gametes (developing and mature) to total area occupied by each of ten acini. This metric, in comparison, provided a measure of the degree to which viable gametes filled the acini located throughout the visceral mass. These metrics were used as indicators of reproductive condition and were compared among organisms located at each of the two study sites using Mood’s Median test. A multiple comparisons test for equal variances was

70 also used to further investigate preliminary observations of gametogenic asynchrony among males and females at Site 1. Further site differences in gametogenic stage, sex ratio, and organism growth (mass), were determined by applying appropriate transformations, when required, and using the two-sample Student’s T-test when all parametric assumptions, including normality, were met. Otherwise, comparisons were made using Mood’s Median, Chi Square or Fisher’s Exact tests (Sokal and Rohlf, 1998). All analyses were conducted using Minitab Software v. 17.2.1, unless otherwise stated.

E. complanata Relocation Experiment

Fifty individuals at each site were collected, massed, maximum valve length recorded, and uniquely marked (valve etch) in October 2010. Organisms were randomly placed into one of two groups at each site (Reference / Experimental) and either returned to the substrate (Reference groups) or placed in coolers and relocated to the alternative study site (Experimental groups).

Attempts were made to resample these individuals between 2011 and 2013, and, for recovered organisms, growth and reproductive development were assessed. One-Way ANOVA was used to compare mass changes among groups and multiple-comparisons were made by Tukey’s all- pairwise approach. Chi-Square tests were used to compare observed gametogenic stages between groups.

Passive Surface Water Sampling

AQUASENSE-P Polar Organic Chemical Integrative Sampling Devices (POCIS) (EST

Laboratories, St. Joseph, MO), consisting of an Oasis HLB solid sorbent, secured between two

71 polyethersulfone (PES) membranes, were deployed seasonally for 60-day sampling periods at each study site. Oasis HLB AQUASENSE-P samplers were employed in this study and align with

USGS Guidelines for the Use of the Semipermeable Membrane Device (SPMD) and the Polar

Organic Chemical Integrative Sampler (POCIS) in Environmental Monitoring Studies (Alvarez,

2010). Oasis HLB has been termed a universal sorbent and has been utilized in the analysis of a wide variety of chemicals in environmental sampling (2010).

POCIS discs were shipped under Argon gas in clean, airtight containers. Containers remained sealed until time of deployment. Upon reaching the deployment site, POCIS discs were transferred to stainless steel deployment canisters (EST Labs) in triplicate, secured to large rocks or concrete blocks, and submerged onto the substrate. A single canister holding 3 POCIS was deployed at each site during each sampling period. Care was taken to ensure sampling discs were oriented perpendicular to water flow to ensure adequate flow across membranes, and were also oriented to ensure that plastic zip-ties were located downstream of the sampling discs as to avoid potential contamination via leaching of organic chemicals that could have been sequestered within the sampler. Field blanks (n = 1 per sampling event per site) were utilized for quality control purposes and in order to ensure that deployment, retrieval, storage, processing, and analysis did not have a significant effect on deployed discs. Blank discs were returned to their original containers and stored frozen during the deployment period.

Upon returning to the laboratory, POCIS discs were gently cleared of debris and disassembled. Solid sorbent from each POCIS was transferred to a glass column packed with clean glass wool and extracted with 40 mL HPLC grade-Methanol (MeOH). Extracts were pooled to make a single, 3-disc composite sample in order to maximize potential analytical response. Extracts

72 were concentrated under a gentle stream of Nitrogen, resuspended in a final volume of 3 mL

MeOH, and stored at -20°C for analysis with the recombinant yeast estrogen screen (YES).

Yeast Estrogen Screen (YES)

POCIS extracts were analyzed using a recombinant yeast estrogen screen (YES) in order to measure the total estrogenic activity of sampler extracts following deployment. This procedure utilizes a recombinant form of Saccharomyces cerevisiae (obtained courtesy David Alvarez, USGS) that has been transfected with the human estrogen receptor, hERα, as well as a plasmid containing the lac-Z reporter gene and estrogen response elements (ERE) (Nelson, 2007). In this assay, the presence of chemical(s) with any affinity for hERα binding elicit a cascade of events that result in β-galactosidase production and an equivalent colorimetric response via β-galactosidase complexation with the chromogenic substrate, chlorophenyl red D-galactopyranoside (CPRG).

While multiple protocols have been developed for this assay (Routledge and Sumpter, 1996;

Rastall et al., 2004), here, all yeast growth and assay media were mixed fresh and syringe filtered

(0.2 µm pore size) immediately prior to use, and a newly purified recombinant yeast culture was transferred from streak plate to growth media and cultured for approximately 24 – 36 hours just prior the start of each assay. All setup was conducted using aseptic technique in order to minimize bacterial contamination of assay components. All tests were conducted using a standard 96-well microplate. A 17β-estradiol positive control was employed in the first row of each test plate, and solvent-only negative controls were utilized between each sample row. Extracts (10 µL extract +

190 µL MeOH or 200 µL extract) were added to the first well of each sample row, and serially diluted across the plate with MeOH (Table 3.1). Plate IDs, number of POCIS per composite, mass of POCIS per composite, maximum %POCIS per test, and extract volume tested are listed in Table

3.2. Each POCIS extract was syringe filtered (0.2 µm pore size) immediately prior to addition to the plate. Plates were allowed to evaporate to dryness before addition of 200 µL assay medium, which consisted of approximately 4x107 recombinant yeast cells, growth media, and CPRG. Each plate was covered, sealed with autoclave tape, and incubated at 30°C until optimal color development in the positive control, typically 24 – 48 hours. Absorbance was measured for each well at 540 nm (colorimetric response) and 620 nm (turbidity) on a BMG Fluostar OPTIMA microplate reader (BMG LABTECH, Germany). One-Way ANOVA was used to compare response data among groups and multiple-comparisons were made by Tukey’s all-pairwise approach.

Elliptio insulsa 17β-Estradiol Exposure Experiment

Acclimation and Setup

In June 2012, approximately 80 E. insulsa were transported to the Columbus Zoo and

Aquarium Freshwater Mussel Conservation and Research Center (Columbus, OH). Organisms were purchased (Carolina Biological Supply, Burlington, NC), shipped overnight under moist conditions, and immediately transferred to continuous-flow artificial raceways upon arrival at the facility (Image 3.5). Raceways were supplied with water pumped directly from the Scioto River and were filled with clean gravel (depth ≈ 4”). Organisms were acclimated and held for approximately four months prior to initiation of experimental procedures in order to ensure survival and adequate organism health following transport and introduction to raceway conditions. Survival/health were assessed by observing adequate valve closure response to an external stimulus (gentle agitation of posterior valve margins). Following the acclimation period

(October 2012 ), organisms were drawn from the substrate, mass (g) was recorded, and a biopsy of gonad tissue was collected. The biopsied tissues were qualitatively analyzed for determination

74 of sex, and sexed individuals were uniquely marked (valve etch). This procedure was repeated until 20 experimental E. insulsa of each sex were identified. These organisms were returned to the raceway substrate and held for an additional two months in order to ensure survival and re- acclimation following the biopsy procedure and in order to initiate experimentation at the expected earliest stages of the subsequent gametogenic cycle.

Experimental Design

Male and female E. insulsa were each randomly assorted into one of four groups (n =

5/group/sex) and were dosed with 10 µL of one of three 17-β estradiol (E2) solutions or a solvent blank. Groups were defined by exposure level; “High Dose” (19 ng/µL E2), “Intermediate Dose”

(1.9 x 10-2 ng/µL E2) or “Low Dose” (1.9 x 10-5 ng/µL E2). Exposure media were made fresh prior to initiation of the exposure by dissolving E2 (Sigma Item #E2758) in 27% EtOH and diluting, as necessary, with 27% EtOH to reach each of the final desired concentrations. At time zero, organisms were collected from raceways and valves were gently pried apart using snap ring pliers and held open by inserting a syringe cap between the valves. Dosing was carried out via direct injection of exposure media (exposure groups) or pure solvent (solvent control) into the gonad.

Each organism was dosed twice (time = 0, time = 5 days) and all individuals were returned to the raceway immediately following each dosing event. All organisms were held in the same raceway throughout the duration of the initial test period (10 days) and for an additional 5 months following exposure. Scioto River water temperature during the 10-day experimental period ranged from 2 – 5 °C and the light/dark cycle was maintained at 12hr/12hr to simulate daylight hours at the time. At time = 10 days, organisms were again pulled from the substrate and a biopsy of gonad tissue was collected, as described previously, with one modification in treatment of male

75 samples, which were not preserved in NBF, in order to allow immediate measurement of observed spermatozoa motility. Organisms were then held in the raceway for an additional 5 months and, at that time, were biopsied again and subsequently sacrificed for histological examination of the gonad.

Female Exposure Histological Analysis

For females, gametogenic stages were histologically determined based upon proportion of the gonad occupied by atretic, developmental, or mature tissue. Acini were selected for analysis according to procedures described in Chapter 2 (see Methods – Histological Processing and Analysis). Areas (µm2) occupied by each of the three tissue types (atretic, developmental, mature) were measured for each acinus selected for analysis and relative proportions were used to stage gametogenesis, as was previously described (see Chapter 2). This process was conducted twice for a total analysis of 10 acini per organism. Proportions of gamete stages observed in histological samples were compared and group differences were statistically determined using

Chi Square. The proportion of atretic tissue present in measured acini was also compared between exposure groups using one-way ANOVA.

An identical approach to acinar selection was utilized for calculation of the AAF. Acini were selected as previously described, and the AAF was calculated as the proportion of area filled with viable gametes (developing and mature) to total area occupied by each of ten acini. AAF values were compared among exposure groups using one-way ANOVA and individual group comparisons were made by Dunnett’s Multiple Comparisons with a Control.

Female Exposure Biopsy Analysis

Female biopsy samples were collected at 10 days post-initial exposure as well as 5 months post-initial exposure in order to assess both acute effects of E2 exposure, as well as any potential chronic, delayed effects on the gametogenic cycle. Initial 10-day samples were analyzed immediately following their collection (unstained) by aliquotting a 10-µL sample directly onto a clean glass slide after mixing to ensure homogeneity of the fluid sample. Samples were imaged under 4x magnification and areas were recorded for all observed ova. Samples collected following the 5-month developmental period post-exposure were treated as was described previously (see

Biopsy Preparation in Methods, Chapter 2) and stained samples were again imaged under 4x magnification and areas were recorded for all observed ova in a 10-µL aliquot. Differences in median ova area values among exposure groups were determined using Mood’s Median Test.

Male Exposure Histological Analysis

Gametogenic stage was assigned to male histological samples according to the procedure presented earlier for male freshwater mussels using Cacapon River E. complanata (see Figure 2.3).

Random selection of acini for analysis was conducted according to the procedures described above for female samples. Effects among exposure groups were compared for gametogenic stage,

% atretic tissue, % mature tissue, and AAF. Categorical data (gametogenic stages) were analyzed by Chi Square. When parametric assumptions were met, groups were compared by one-way

ANOVA and individual differences between groups were tested by Dunnett’s Multiple

Comparisons with a Control. Otherwise, group responses were compared using Mood’s Median

Test.

Male Exposure Biopsy Analysis

Male biopsies were prepared and analyzed according to previous descriptions (see Male

Biopsy Tissue Analysis in Methods – Chapter 2) and biopsy-based gametogenic stage was classified according to the procedure developed for freshwater mussels using Cacapon River E. complanata

(see Figure 2.8). Biopsy-based and histologically determined gametogenic stage designations were compared using Attribute Agreement Analysis and agreement was assessed using Cohen’s

Kappa statistic in order to validate the procedure for this unionid species. Differences in biopsy- based gametogenic stage designations were assessed using the Chi Square test. Proportion of mature tissue present in biopsies of individuals among exposure groups was compared using one- way ANOVA and individual differences between groups were tested by Dunnett’s Multiple

Comparisons with a Control.

Results

Preliminary Sampling and Histological Analysis

Qualitative (gametogenesis) and quantitative (gender frequencies) histological analysis of

E. complanata (n = 19/site) sampled from each of four sites located within upper, mid, and lower reaches of the Cacapon River suggested sex ratios skewed toward females (2.8:1) and highly variable gametogenic development in females sampled at the uppermost site sampled (Site 1), while at the most downstream site (Site 2), female gametogenic stages were observed to be

78 consistent among all females sampled (Figure 3.1) and sex ratio, while still skewed toward females

(1.7:1), was closer to a predicted ratio of 1:1 for unionid mussels (Dillon, 2004). Subsequent biopsy sampling during the 2009 season (nSite 1 = 19; nSite 2 = 23) supported initially calculated differences between these two sites, as significant departure from the expected 1:1 ratio was observed for

Site 1 (Chi Square p = 0.0012, α = 0.05) but not for Site 2 (Chi Square p = 0.22, α = 0.05).

Morphometrics and Sex Ratios

A total of 263 E. complanata were sampled between September 2009 and October 2013

(n site1 = 129; n site 2 = 134). Initial mass values recorded for sampled individuals (x ̅ site 1 = 64.3 g, SD

= 22.4 g; x ̅ site 2 = 43.6 g, SD = 12.8 g), as well as maximum valve length (x ̅ site 1 = 82.4 mm, SD = 9.7 mm; x ̅ site 2 = 74.4 mm, SD = 7.4 mm) differed significantly between sites (two-sample Student’s T-

Tests: p < 0.001, α = 0.05). These differences are depicted in E. complanata mass (Figure 3.2) and maximum valve length distributions (Figure 3.3). Sex was positively identified for 187 of the organisms sampled (n site 1 = 85; n site 2 = 102) and no sex-dependent differences in either mass or length were detected at either site (Table 3.3). Of the 187 sexed organisms, 87 males and 100 females were observed. Site specific gender differences are presented in Table 3.4. Though the

Site 1 population exhibited a greater departure from the expected ratio than the Site 2 population

(Figure 3.4), no significant departure from an expected 1:1 ratio was detected for either Site 1 or

Site 2 (Chi-Square Goodness-of-Fit: p site 1 = 0.233; p site 2 = 0.843, α = 0.05).

Gametogenic Assessments

Of the organisms sampled in May 2013, five males and five females were observed per site. Seven males, three females and six males, four females were observed in the July/August sampling at Sites 1 and 2, respectively. Gametogenic stages observed for both male and female

E. complanata sampled in May 2013 are shown in Figure 3.5. Site 1 males exhibited a range of developmental stages including Late Active, Mature, and Resorbing, while Site 2 males generally exhibited Late Active development, though one individual was classified Mature. No significant differences in male staging were detected between sites when the single Resorbing individual from Site 1 was omitted from analysis (Fisher’s Exact Test p = 0.343, α= 0.05). Additionally, no significant atypical spermatogenesis was observed in May samples. All Site 2 females sampled in

May were classified as Mature, while those at Site 1 exhibited characteristics of Developing,

Mature, and Resorbing gametogenic stages. These differences observed for female gametogenic stages between sites in May were found to be significant (Fisher’s Exact Test p = 0.048, α = 0.05), however, due to the limited number of individuals sampled, only Mature and Not Mature

(category includes both developing and resorbing staged individuals) classifications were considered for this comparison. All females collected and sampled in May at Site 2 were classified

Mature while 80% (4 of 5) were classified Not Mature at Site 1. Developmental classifications appeared to have undergone a significant shift as of the July/August sampling, as all males at each site exhibited atypical spermatogenesis and females were observed to be undergoing gonad resorption (Figure 3.6). No between site differences were observed for either sex among

July/August samples.

All organisms sampled and staged were further categorized into one of two groups

(“Mature”, “Not Mature”) for analysis of gametogenic synchrony between sexes at each site. The

80 only asynchrony detected was at Site 2 among individuals sampled in May, as all females were categorized as “Mature” while four of five males previously staged Late Active were considered

“Not Mature” (Fisher’s Exact Test p = 0.048, α = 0.05). No further differences in degree of gonad maturation were detected between male and female E. complanata at either site.

AAF/GAF

The AAF and GAF were calculated for each organism sampled for histological analysis in

2013 (Spring and Summer sampling events). Median values for each metric were compared between sites for both sexes during each sampling period and distributions are represented in

Figures 3.7 (AAF) and 3.8 (GAF). No between-site differences in either metric were detected, regardless of gender or sampling date (Mood’s Median tests, all p values > 0.05). However, observed distributions indicated highly variable metric scores for both male and female E. complanata at Site 1, relative to scores observed at Site 2. This observation was statistically evaluated by multiple comparisons test for equal variances for Spring AAF and GAF samples of both sexes and significance was detected for male E. complanata between sites for both AAF (pmale

= 0.022, α = 0.05; pfemale = 0.065, α = 0.05) and GAF (pmale = 0.008, α = 0.05; pfemale = 0.136, α =

0.05). Median values of AAF and GAF for females each dropped significantly between Spring

(Median AAF = 73.7; Median GAF = 68.9) and Summer (Median AAF = 0; Median GAF = 0) sampling events (Mood’s Median Test pAAF = 0.001, pGAF = 0.001, α= 0.05). A similar, though less drastic, decrease was observed for male GAF between Spring (Median = 83.8) and Summer (Median =

58.4) sampling (Mood’s Median Test p = 0.005, α = 0.05) though no difference was observed for male AAF between these dates (Mood’s Median Test p = 0.835; Spring Median AAF = 86.0;

Summer Median GAF = 85.6).

E. complanata Relocation Experiment

Twenty-month recovery rates for organisms transported from Site 1 to Site 2 (Group

Exp1) and from Site 2 to Site 1 (Group Exp2) were 48% and 52%, respectively, while rates for reference groups at each site were 88% (Site 2 reference organisms = Group R2) and 56% (Site 1 reference organisms = Group R1). Recovery efforts declined following the 20-month sampling event (data not shown). Figure 3.9 depicts average mass change and 95% CIs for each group at time = 20 months. Because significant differences were detected for organism mass between sites, group differences in 20-month mass change (g) were compared using the General Linear

Model function in Minitab with initial mass (g) as a covariate and Tukey’s simultaneous tests for differences of means was used for individual group comparisons. After accounting for initial mass, significant differences in 20-month mass change were detected among groups (Figure 3.10), as

Tukey’s pairwise comparison approach detected significant differences between groups Exp1 and

Exp 2 (p = 0.001, α = 0.05), and between groups R2 and Exp2 (p = 0.001, α= 0.05). While Exp1 organisms (x ̅ = 1.02 g; SD = 1.80 g) exhibited greater mass increases than controls (R1), these differences were not found to be significant, and no differences could be detected for mass changes between R1 and Exp2 groups.

Gametogenesis was evaluated during 2013 sampling when conditions allowed for access to experimental organisms. River flow conditions in May 2013 prevented access to experimental organisms at Site 2. Relocated individuals from group Exp2 were located at Site 1 in May (n = 6 male; n = 5 female), however no organisms from the R1 group were recollected during this sampling event for comparison. Group Exp2 males in May were designated as either Late Active

(n = 4) or Mature (n = 2) based upon biopsy analysis, while female classifications ranged from

Spent/Resorbing (n = 3) to Developing (n = 1) and Mature (n =1). July sampling resulted in

82 recollection of nine Exp1 organisms at site 2, four Exp2 organisms at Site 1, and one organism each from groups R1 and R2. All Exp2 organisms identified in this sample were female and all were classified as Spent/Resorbing (n = 4). The single R1 control was also found to be a Spent/Resorbing female. Of the nine Exp1 organisms collected at Site 2 in July, five males ranged in gametogenic classification from Spent/Resorbing (n = 2) to Early Atypical (n = 1) as well as Late Atypical (n = 2).

The remaining Exp1 organisms were all Spent/Resorbing females (n = 4). The single R2 control was identified as a Spent/Resorbing Female. The final sampling (October 2013) resulted in successful recollection of experimental organisms only at Site 2 (nExp1 = 7; nR2 = 7). Of those sampled, one

Exp1 male was identified as Spent/Resorbing while five R2 males were collected and classified as either Mature (n = 4) or Spent/Resorbing (n = 1). Two R2 females were observed in this sample

(one Mature and one Spent/Resorbing), while six Exp1 females were identified, and ranged in gametogenic stage classification from Spent/Resorbing (n = 3) to Developing (n = 1) and Mature

(n = 2). Site comparisons of seasonal gametogenesis were not possible due to low recollection frequencies experienced for at least one of the two sites during each sampling event.

POCIS and Extract Estrogenicity

Samplers (3 POCIS/site/deployment) were successfully deployed and recovered at each site and compared during Summer/Fall 2011, Spring/Summer 2012, and Summer/Fall 2013.

Deployment periods for each of the three sampling events were 45, 60, and 60 days, respectively.

Concentrated MeOH extracts of composite (3-disc) POCIS samples were tested for total estrogenicity and Estradiol equivalent factors (EEQ) were calculated for each sample using

Equation 1, modified from Alvarez et al. (2008). E2 positive controls were included on each test plate and produced expected sigmoidal absorbance curves when plotted against log transformed

E2 concentrations (M) (Figure 3.11). Positive control EC50 values, or the concentration (M) of E2 producing 50% of the maximum E2 colorimetric response, were variable between plates (Table

3.5), therefore calculations for each test plate were made using the positive control data from that plate in place of average E2 response data. EC20 values were used in place of EC50 for the calculation of EEQ, in order to minimize predictions beyond measured sample responses as sample extracts generally failed to produce complete sigmoidal response curves and did not generate absorbance values equivalent to EC50 for E2 standards (Jarosova et al., 2012).

Equation 1. EEQ (ng E2/POCIS) = E2 EC20 (ng E2/mL) / Test EC20 (POCIS/mL)

Maximal response (corrected absorbance) for POCIS extracts tested corresponded with E2 equivalent concentrations (M) from < 7.33E-12 (LOD) at Site 2 in June 2012 to 1.63E-09 at Site 1 in October 2011 (Table 3.6). Season/site differences are displayed in Figure 3.12.

Positive EEQ values were determined for two of the six POCIS samples, Site 1, October

2011 and Site 1, June 2012 (0.22 ng/POCIS and 1.26 ng E2/POCIS, respectively). One sample (Site

2, June 2012) exhibited no significant response above negative (solvent) control, resulting in an

EEQ of zero. Due to the limited ability to determine EEQ values for POCIS extracts, site differences for each deployment period were assessed using maximal response E2 equivalent (M) concentrations (Figure 3.12). Significant differences were detected among the maximal response

E2 equivalent values observed (one-way ANOVA; p = <0.001, α= 0.05). Significant differences between Site 1 and Site 2 values were detected for both the October 2011 and June 2012 deployments (Tukey’s Pairwise Comparisons; 95% Confidence), while no difference was detected for 2013 samples. Trip blanks (one POCIS/site/deployment) exhibited no responses above negative controls.

Elliptio insulsa 17β-Estradiol Exposure

10-Day Male Gamete Effects

Tissues collected in biopsy samples from male E. insulsa at time = 10 days were immediately transferred to a Neubauer hemocytometer and motility of spermatozoa was manually assessed. As methods proved highly inaccurate, no viable results for 10-day effects in male organisms were obtained.

10-Day Ova Area Effects

Viable ova were extracted via nonlethal biopsy 10 days post the initial dosing event from all female organisms in Control (n = 5), Low (n = 4), and Intermediate (n = 4) dose groups (Images

3.6 – 3.8). One individual in the Low dose group was found to have suffered a partially crushed valve early in the exposure period and did not survive, while one individual in the Intermediate dose group had been misidentified initially as female and was found to be male following experimentation. High dose organisms (n = 5) exhibited atrophy of all observed oocytes in three of five exposed organisms (Image 3.9) and no viable ova were available for comparison in biopsies collected form these individuals. Biopsy samples from the final two exposed organisms in the high dose group displayed viable ova characteristic of late development. When ova areas (µm2) were pooled from all organisms within each exposure group (Figure 3.13), significant differences in median ova areas were detected (Mood’s Median Test, p = 0.002, α = 0.05). These differences were detected between exposure groups, as the Low dose group (median area = 9901 µm2) exhibited significantly lower median ova areas than either the Intermediate (median area = 12820

µm2) or High (median area = 12820 µm2) dose groups. No differences were detected for any individual exposure group when compared with solvent control (median area = 11314 µm2).

Five-Month Female Effects

Organism survival was 100% in all groups during the five months following exposure.

Following this period of potential gametogenic activity (May, 2014), organisms were pulled from the raceway substrate, biopsied, and immediately sacrificed by separation of both anterior and posterior adductors and excision of the foot/visceral mass complex. Excised tissues were immediately immersed in a pre-labeled 50-mL centrifuge tube filled with 40 mL NBF in order to preserve tissues for histological processing.

No significant differences among groups were observed for female AAF values observed in histological sections 5 months post-exposure (one-way ANOVA, p = 0.130, α = 0.05). Exposure group versus control comparisons were made using Dunnett Multiple Comparisons with a Control and results are listed in Table 3.7. Overall, AAF values increased with increasing dose, however these increases were not significant. Average AAF values ranged from 59.1 (Low dose) to 65.5

(High Dose), while control values averaged 60.6 (Figure 3.14).

Viable ova were again extracted via nonlethal biopsy 10 days post the initial dosing event from all female organisms in Control (n = 5), Low (n = 4), Intermediate (n = 4) and High (n = 5) dose groups (Images 3.10 – 3.13). A mixture of developing, mature, and atretic ova were observed for all exposure levels. When ova areas (µm2) were pooled from all organisms within each exposure group, no significant differences in median ova areas were detected (Mood’s Median Test, p =

0.057, α = 0.05). The Low dose group exhibited the lowest median value observed in the 5-month biopsy sampling (median area = 7958 µm2), while the Intermediate dose group exhibited the

86 highest median ova area (9230 µm2). Ova areas observed in High dose and control organisms fell between these values, each having a median value of 8582 µm2 (Figure 3.15).

No differences were observed for gametogenic staging of histological samples, as all individuals were classified as Developing at the 5-month sampling (all samples identical; Chi

Square could not be calculated). Due to the observation of what appeared as an increased rate of atretic ova in 10 day biopsy samples from the High dose group, the proportion of atretic tissue among analyzed acini was compared, but no significant differences were found for any group, relative to controls (Figure 3.16 – Dunnett Multiple Comparisons with a Control). Two outlier values were omitted from the analysis (43.77 % - Intermediate Dose; 40.45 % - Control) in order to meet the parametric assumption of normality. The only significance was observed between the

Low dose (x ̅ = 17.05 %, SD = 3.90%) and Intermediate dose (x ̅ = 5.31 %, SD = 2.26%) groups (one- way ANOVA, p = 0.014, α= 0.05).

Male Staging Procedure Validation

The procedures for gametogenic staging of male freshwater mussels presented in Figures

2.3 (histological approach) and 2.8 (biopsy-based approach) were employed for male E. insulsa histological and biopsy samples in order to validate the methodology for this Elliptio species as well as to support the utility of biopsy data analysis in addition to traditional histological assessment for response to E2 exposure. Gametogenic stage determinations were made for each individual included in the study (n = 20) using the alternative approaches and results are presented in Figure 3.17. Late Active and Mature stages were generally observed, and two biopsy samples failed to correspond correctly with the histologically-based stage determination. One individual was incorrectly designated Late Atypical by biopsy analysis, as the % atypical cells

87 observed (1.51%) was slightly above the defined threshold for Atypical categorization (Atypical

Classification Threshold ≥ 1.5%) and Attribute Agreement Analysis resulted in a Kappa score of

0.80, which suggests “Substantial” agreement (see Table 2.4) between the two methods.

Five-Month Male Effects

No significant differences between exposure groups were detected for either histology- based or biopsy-based (single Late Atypical sample omitted) gametogenic stages assigned (Chi

Square phistology = 1.000; pbiopsy = 0.802 α = 0.05). Average AAF values for each exposure level (n =

5 organisms per exposure level) were found to be significantly lower than controls (one-way

ANOVA, p = 0.005, α = 0.03) and these differences (Figure 3.18) were observed between control and exposed organisms, regardless of exposure level ( xCTRL̅ = 85.5, SD = 2.4; xLOW̅ = 79.0, SD = 3.0; xINT̅ = 79.3, SD = 2.7; xHIGH̅ = 80.5, SD = 1.8). Exposed organisms also exhibited significant differences from controls for the percentage of atretic tissue occupying measured acini (Figure 3.19). This difference was not detected in biopsy tissue (Mood’s Median Test p = 0.261, α= 0.05), though both Intermediate and High dose groups exhibited a broader range of values than either Low dose or controls (Figure 3.20).

The average proportion of mature tissue present in measured acini, as well as the proportion of mature tissue present in biopsy aliquots sampled, exhibited no differences among groups and no differences between control and any exposure level (phistology = 0.602 pbiopsy = 0.372;

α= 0.05).

Discussion

Investigation into potential endocrine disruption or other reproductive impairment in E. complanata residing in the Cacapon River was undertaken following the observation of such effects in fish, namely smallmouth bass, Micropterus dolomiue (Blazer et al., 2007). Preliminary sampling suggested the potential for sex ratios skewed toward female differentiation and asynchronous gamete development among males and females of, particularly, one population of

E. complanata located near the town of Wardensville, WV (Site 1). This site is located in the upper reaches of the Cacapon River, and lies in close proximity to an upstream region of the watershed characterized by a high concentration of independent poultry rearing facilities (Elrashidi et al.,

2008). Litter waste produced at these facilities is most often composted and applied locally within the basin as a soil amendment or as a waste disposal method (Chambers and Leiker, 2006). Several xenobiotics of concern have been associated with application of poultry litter, including, among others, the steroid hormone, E2 (Nichols et al., 1997). Controlled laboratory studies on the persistence of E2 and its primary metabolite, estrone (E1) in soils seem to support rapid degradation and efficient biotic transformation to highly sorbed, non-bioavailable products,

(Colucci et al., 2001). However, as is discussed extensively in a review of E2 in the environment by

Khanal et al., (2006), field studies following land application of manure and poultry litter indicate significant potential for environmental transport and contamination of aquatic systems, as E2 concentrations have been observed in runoff at 3500 ng/L (Nichols et al,. 1998) and in spring waters up to 66 ng/L (Peterson et al., 2000). Efforts were therefore made to investigate any potential correlations between reproductive impairment in E. complanata and surface water estrogenicity in the Cacapon River. This was done by locating a suitable “reference” site for

89 comparative purposes (Site 2). Site 2 was selected as a “reference” as it is located in the far lower reaches of the watershed in an area of dense forest and very little agricultural or industrial activity

(Constantz et al., 2005). Subsequent sampling detected no significant differences among organisms within each of these populations for sex ratio or seasonal gametogenesis. However, significant between-site differences were observed for AAF and GAF variance in spring male E. complanata samples. Site 1 males exhibited significantly greater variance when standard deviations of measured values were compared. While no evidence was detected to suggest differences in assigned gametogenic stages among organisms, these metrics, which provide measures of reproductive condition, suggest differences in gametogenic development for males between the two sites. Females demonstrated similar patterns of AAF and GAF, however differences were not found to be significant.

No evidence of intersex was observed at either site, as it had been previously reported at high incidence rates for fish in the region (Blazer et al., 2007). The concept of intersex, or development of female tissue within a predominantly male gonad (and less commonly male tissue in the female gonad) in bivalves, has been referenced as both a direct effect of exposure to environmental contaminants (Chesman and Langston, 2006; Ortiz-Zarragoitia and Cajaraville,

2010), as well as a “curious” physiological occurrence (Heard, 1979). E. complanata has been previously described as an “occasional hermaphrodite,” (1979) however the hypothesis that a number of dioecious unionid species exhibit some degree of intersex condition as a normal physiological response to their environment has not been sufficiently explored or supported, particularly in relation to the potential role that environmental contaminants, such as steroidal hormones and xenobiotic hormone mimics, might play in the incidence of this condition. Here, low levels of detectable surface water estrogenicity in extracts collected from POCIS samplers did not correspond with any observed incidence of intersex condition in E. complanata, nor did the

Site 1 proximity to known nonpoint sources (field application of poultry litter and other agricultural activities) of potential surface water contamination.

Gametogenesis in Cacapon River E. complanata appeared to follow a seasonal pattern of spring/early summer maturation and subsequent spawn, followed by resorption (female) of retained tissue or atypical development (male) in the following months, based upon both histological and biopsy analyses. This finding is similar to previous reports on the life history of this species in other regions (Matteson, 1948). Elucidation of a particular functional role for atypical spermatogenesis in unionid mussels has yet to be fully accomplished in the literature, though a number of hypotheses for the potential role of this gametogenic pathway have been proposed. In a detailed structural study of atypical development, Shepardson et al. (2012) discuss a number of these hypotheses, and conclude that a functional reproductive role of atypical spermatozoa in unionids is most likely, and is possibly associated with maintenance of dioecy and equal sex ratios within unionid populations. These organisms have been shown to possess the unique characteristic of high fidelity doubly uniparental inheritance of mitochondrial DNA (DUI)

(Breton et al., 2007). According to Shepardson et al., (2012), maintenance of two separate spermatogenic pathways may lend to the ability of male organisms to selectively pass gametes having either female (F-type) or male (M-type) mitochondria, which could, in turn, play a role in gender development in resulting offspring, thus providing a mechanism by which equal sex ratios could be maintained. Both typical and atypical (to a much lesser extent) spermatogenesis was observed in this study during predicted periods of active gametogenesis in spring, which supports the hypothesis of an active reproductive role for atypical spermatozoa. However, the observance of a high degree of atypical spermatogenesis in late summer and autumn sampling, corresponding with high rates of gonad atrophy in females, possibly suggests a secondary role of the atypical

91 pathway, though the function of any secondary role played by this tissue will not be speculated herein.

Comparative POCIS sampler data were limited in this study to one spring/early summer

(2012) deployment and two late summer/early autumn deployments in 2011 and 2013. Additional deployments failed to provide useful data due to vandalized or otherwise lost samples, which is an important consideration in decisions to utilize these passive sampling devices (Alvarez, 2010).

While the estrogenic potential of surface waters in the Cacapon River appears to be present at both sites, positive YES assay responses for POCIS sampler extracts, when detected, were generally weak. Site 1, however, did appear to generally exhibit a higher degree of estrogenic activity than the reference, Site 2, as was predicted due to the potential influx of chemicals associated with agricultural activity in the area. It is important to point out that these responses are indicative of the presence of some chemical or chemicals that have the ability to positively interact with the vertebrate estrogen receptor. The identification of chemical(s) contributing to this response, however, cannot be speculated and is beyond the scope of this work. Possibly as important was the observance of either apparent toxicity or some non-toxic inhibition of yeast activity in assayed extracts, particularly at the highest concentrations tested for both October

2013 samples, as well as the October 2011 (Site 2) sample. Similar findings have been reported in the literature (Alvarez and Perkins, 2006). The possibility exists that this response is an artifact of sample extraction associated with the presence of residual compounds from manufacture of the

POCIS in the sorbent matrix that co-elute with those compounds accumulated during deployment or, alternatively, an effect resulting from sample processing. Positive controls (E2) did not exhibit this type of response, so contamination associated with solvent or assay medium can, at minimum, be ruled out. Trip blanks failed to exhibit any response above background, so the potential effect of toxic compounds on yeast activity in those samples could not be determined.

Therefore, the potential of these apparent toxic or inhibitory effects to be associated with chemicals accumulated during POCIS deployment in Cacapon River surface waters cannot be either supported or refuted at this time.

While the focus of this study, as has been the case to date for most investigations into possible endocrine disruptive potential of surface waters, was associated with estrogenic effects of natural and synthetic estrogens in the environment, many compounds exist that either mimic androgenic activity or inhibit observed estrogenic responses of samples that, in their absence, could exhibit significantly greater estrogenic potential (Yang et al., 2015). Utility of the YES assay for quantifying antiestrogen activity has been demonstrated in the literature (Buckley, 2010) and requires little in terms of resources or efforts outside those required for typical YES analysis.

Wastewater treatment plant (WWTP) effluent samples tested in the development of this analytical approach indicate as much as a 50% reduction in estrogenic response attributed to antiestrogenic activity in samples tested (2010). Additionally, the presence of commonly used chloro-s-triazine herbicides, such as atrazine, have been shown to inhibit the estrogenic activity of some estrogen mimics up to 66% (Rastall, 2004). As environmental samples often contain a complex mixture of chemicals across a range of concentrations, each potentially having a unique interaction potential with the estrogen receptor utilized in this assay (ERα), analysis of both estrogenic and antiestrogenic activity is recommended for any future investigation into the estrogenic potency of environmental samples. It is unknown whether estrogenic responses observed in this study were in any way influenced by antiestrogenic potency of sample extracts.

Organisms were relocated between sites 1 and 2 in order to test the hypothesis that observed differences in growth and gametogenesis were site dependent and reversible. Organism growth, as measured by mass changes, suggest that organisms at Site 1 are impacted by some

93 factor or combination of factors that significantly influences growth or seasonal changes in total wet weight, as significant decreases were detected between reference site controls (R2) and individuals relocated from Site 2 to Site 1 (Exp 2), while those same relocated individuals exhibited no difference in mass changes when compared with Site 1 controls (R1). Alternatively, organisms moved from Site 1 to Site 2 (Exp1) appeared to experience positive mass changes in comparison with Site 1 controls (R1), although significance was not detected for this increase. The Exp1 group, however, did experience significantly increased mass in comparison to Exp 2 organisms. This is an interesting observation, as initial mass for organisms originally found at Site 1 was significantly higher than that for Site 2 organisms, and, a deceleration of growth is to be expected with increasing mussel size (Anthony et al., 2001). Environmental conditions, however, are known to play a role in limiting the ability of mussels to maximize growth rates (Bauer, 1992) and, therefore, the observation of site-dependent effects on organism growth is not altogether unexpected.

Kesler et al. (2007) reported similar observations in E. complanata transported between three lakes in Rhode Island, and in that study attributed growth cessation and low condition indices measured to food limitation, which is a potential factor that cannot be discounted in this case.

Seasonal gametogenesis was also evaluated for relocated organisms using the nonlethal biopsy technique. Severely limited sample sizes for recollected individuals during each sampling event prevented between-site comparisons. Qualitative analysis of stages observed suggests possible gametogenic asynchrony among females even at Site 2, although these observations were made for organisms that had been relocated from Site 1, where the initial histological observations of gametogenic asynchrony among females were observed. While Site 1 organisms tended to exhibit a higher rate of Spent/Resorbing classification during periods of expected active gametogenesis – particularly in spring and particularly among females sampled – in addition to greater variation in stages exhibited, the limited sample sizes in this study and the potential

94 variability inherent to biological samples limits the ability to relate any of these observations to some factor or factors associated with their environment.

Females at Site 1 did appear to exhibit a degree of gametogenic asynchrony in comparison to Site 2 females, and males at Site 1 exhibited significantly higher degrees of variance in reproductive condition (AAF) among individuals sampled relative to males at Site 2. While estrogenic responses associated with POCIS extracts were generally weak, Site 1 extracts also exhibited increased total estrogenicity in comparison to Site 2 for multiple deployments. These data support the original hypothesis of some correlation between surface water estrogenicity and reproductive effects in E. complanata in the Cacapon River. However, available comparisons in this study were limited due to a variety of factors, including limited sample sizes, low recovery of experimental organisms, high water levels preventing organism access, and sampling units that were vandalized or otherwise lost during deployment.

To further investigate a possible relationship between exposure to vertebrate estrogens and reproductive effects in freshwater mussels, an exposure experiment was conducted using the unionid mussel, E. insulsa. Organisms were exposed to one of three nominal E2 concentrations

(19 ng/µL, 1.9 x 10-2 ng/µL or 1.9 x 10-5 ng/µL E2) via direct injection of exposure medium to the gonad and effects on gamete development were assessed at 10 days, as well as five months, post- exposure. Attempts were made to assess motility of spermatozoa observed in male biopsy samples collected 10 days following the start of the experiment and, therefore, those samples were not preserved according to stated methods. However, efforts to manually assess sperm motility were unsuccessful, as methods proved highly inaccurate (data not shown). Female biopsy samples at 10 days exhibited signs of atresia and cell resorption, however viable ova were present in some samples. Organisms exposed to the highest E2 concentration, however, exhibited

95 advanced resorption of ova in three of five organisms tested, and no viable ova were observed in these individuals. The observation was made, however, that several highly enlarged atretic ova had developed in these organisms, though, as these were not considered viable, they were not included in analyses. This observation is similar to findings by Wang and Croll (2004) who observed occasional greatly enlarged oocytes in the acini of juvenile scallops, though these organisms were exposed to significantly higher concentrations of estradiol (30 µL of 1000 ng/µL solution injected equal to approx. 110 nmol E2 per injection) than were utilized in this study (10 µL of 19 ng/µL E2 at highest dosing level equal to approx. 0.70 nmol E2 per injection). In addition, the authors of that study (2004) compared findings for the five largest ova observed in each sample, while median ova areas were compared here. In this study, for the viable ova observed in the remaining two individuals at the highest exposure level, no difference in median area could be detected from controls, nor were differences observed for median areas among the other exposure groups.

When ova areas were re-examined following 5 months of subsequent development, an increasing trend was observed for ova areas with increasing dose, however no significant differences could be detected between any two groups when median ova areas were compared. Whether significant differences could be detected for the largest five ova in each sample, as was observed in the Wang and Croll (2004) study, was not investigated and is unknown. The lack of detectable response to female gamete development and maturation observed here, though, is in agreement with Scott’s (2013) critical review of the perceived role that vertebrate steroids play in mollusc reproduction and reproductive development. In that review, the argument is constructed that, though a number of studies have positively associated reproductive effects in molluscs to vertebrate steroid exposure, inherent flaws in design and implementation of these studies renders findings insignificant. Additionally, an apparent lack of sufficient evidence for the complete biochemical pathways in molluscs required for biosynthesis of vertebrate steroids

96 suggest that observed instances of vertebrate estrogens in molluscs are more likely a result of exogenous steroid uptake from the environment, as many molluscs have demonstrated the ability to absorb and retain the vertebrate steroids E2 and testosterone (T) by esterification, or fatty acid conjugation (2013), including bivalves (Labadie et al., 2007).

In contrast to female findings, analysis of male tissues, both histologically and biopsy- based, supported some level of biological effect related to organism E2 exposure. While a typical sigmoidal dose-response for E2 effects was not observed for any measured endpoint exhibiting significant departure from controls, this study was designed in order to detect presence or absence of biological effect over a broad range of exposure doses, and, therefore, it is not entirely unexpected that the typical dose-response was not observed for these effects. It is interesting, however, that effects on male AAF were observed in males five months after the exposure regardless of exposure group. As was discussed earlier, previous literature regarding the presence and behavior of vertebrate steroids in molluscs, including bivalves (see Alexander, 2013 and

Labadie at al., 2007) suggests that a majority of injected E2 is likely rapidly conjugated by fatty acids into an esterified form. The biological significance of this mechanism is unclear, however, it is hypothesized that this mechanism of rapid E2 conjugation in molluscs may result in, overall, no significant difference in realized E2 exposure between groups dosed across some concentration ranges, as the levels of active, free E2 resulting from the activity of this mechanism could be rendered, essentially, equal. This type of effect has been observed for the mussel, Mytilus galloprovincialis following exposure of organisms to 20, 200, and 2000 ng/L E2, as the levels of free E2 measured were only observed to increase at the highest dose (Janer et al., 2005). In addition, while vertebrate conjugation of steroids such as E2 results in the production of excretable, polar conjugates, the esterified conjugates formed in molluscs are nonpolar and would more likely be retained in lipophilic tissues (Giusti and Joaquim-Justo, 2013). This is also

97 supported by findings by Janer et al. (2005) as total (free + esterified) E2 exhibited a dose- dependent increase from 2 ng/g (whole tissue) in control organisms to 258 ng/g in the highest exposure group. Histological analyses in that study, as well as in a replicated experiment by the same authors, found accelerated gametogenesis in M. galloprovincialis, however this effect was only observed at the lowest exposure level, and no gametogenic differences from controls were detected for the 200 and 2000 ng/L groups (2005), possibly indicating some threshold value, above which effects are masked or otherwise inhibited. Regardless, in the present study, male organisms exhibited a decrease in gamete production following exposure to E2, as measured by the AAF metric, and no dose-dependent influence on this effect was observed.

While the potential role that vertebrate steroids play in mollusc and bivalve physiology is not yet clear, seasonal changes in detected levels of these hormones, relating to gametogenic stage, have been observed for a number of mollusc species (Giusti and Joaquim-Justo, 2013), which provides support for some active reproductive role of vertebrate or vertebrate-like steroids in these organisms. While effects following injected E2 exposure were not detected for females in this study, an apparent decrease in overall gamete production was observed for male E. insulsa, regardless of dose. Similar observations were made for E. complanata in the Cacapon River, as male AAF values (as well as GAF values not measured for E. insulsa) exhibited increased variance among Site 1 organisms, with ranges indicating depressed gamete development among some individuals within that population, relative to Site 2 mussels. As slight, but significant increases in estrogenic potency of Site 1 surface waters were detected during multiple POCIS deployments in comparison to Site 2, findings of E2 exposures with E. insulsa and field observations for E. complanata appear to agree, supporting the correlation between estrogenic exposure and reproductive impairment in these organisms.

Image 3.1 – Aereal photograph of the Cacapon River at Wardensville, WV (modified from

Constantz et al, 2003. Portrait of a River – The Ecological Baseline of the Cacapon River.) Site 1 is located in the bottom-left of the image, just upstream of the town’s sewage settling ponds.

Image 3.2 – Photograph taken looking upstream in the Cacapon River at Site 1. All organisms were sampled along the left descending bank in the area just above the riffle seen in this image.

100

Image 3.3– Aereal photograph of the Cacapon River near Great Cacapon, WV (modified from

Constantz et al, 2003. Portrait of a River – The Ecological Baseline of the Cacapon River.) Site 2 is located near the center of the image, as the river winds toward the mouth.

101

Image 3.4– Photograph taken looking downstream from a low-water bridge crossing the

Cacapon River at Site 2. All organisms were sampled near midstream just below the riffle seen in this image.

102

Sample Well % POCIS (2011, 2012 extracts) % POCIS (2013 extracts)

1 0.330 6.67

2 1.65E-01 3.34

3 8.25E-02 1.67

4 4.13E-02 8.34E-01

5 2.06E-02 4.17E-01

6 1.03E-02 2.08E-01

7 5.16E-03 1.04E-01

8 2.58E-03 5.21E-02

9 1.29E-03 2.61E-02

10 6.45E-04 1.30E-02

11 3.22E-04 6.51E-03

12 1.61E-04 3.26E-03

Table 3.1 – YES assay row compositions of POCIS sampler extracts assayed. % POCIS represents the maximum percent of the total extracted sorbent mass assayed during each test.

103

Composite Max % POCIS YES Plate ID mg POCIS/composite Extract Tested (µL) # POCIS Assayed

Site 1 2011 3 300 0.33 10

Site 2 2011 3 300 0.33 10

Site 1 2012 3 300 0.33 10

Site 2 2012 3 300 0.33 10

Site 1 2013 3 300 6.67 200

Site 2 2013 3 300 6.67 200

Table 3.2 – POCIS extract characteristics for YES assay testing.

104

Image 3.5 – Raceways located at the Freshwater Mussel Conservation and Research Center in

Shawnee Hills, OH. Organisms were held in the second raceway from the top and exposure groups were kept separated using plastic grates (seen in image).

105

Figure 3.1 – Gametogenic stages initially identified following visual inspection of histological samples for male (nSite 1 = 5, nSite 2 = 7; left) and female (nSite 1 = 14, nSite 2 = 12; right) E. complanata from two sites in the Cacapon River, WV.

106

Site 1 Site 2 18

v 12

b 10

e 8

P 6

0 15 30 45 60 75 90 105 120 15 30 45 60 75 90 105 120 Initial Mass (g)

Panel variable: Site of Origin

Figure 3.2 – Initial mass (g) distributions for E. complanata from each of the two study sites

107

Site 1 Site 2 20

t 10

0 52.5 60.0 67.5 75.0 82.5 90.0 97.5 105.0 52.5 60.0 67.5 75.0 82.5 90.0 97.5 105.0 Initial Length (mm)

Panel variable: Site of Origin

Figure 3.3 - – Initial length (mm) distributions for E. complanata from each of the two study sites

108

Measured p – Value p – Value Location Sex Average (SD) Variable Male, Female Site 1, Site 2

Male 64.7 (19.0) Site 1 0.830 Female 65.8 (22.3) Mass (g) <0.001 Male 43.9 (12.2) Site 2 0.931 Female 43.7 (12.5)

Male 82.9 (8.5) Site 1 0.922 Valve Length Female 82.7 (7.8) <0.001 (mm) Male 74.5 (6.8) Site 2 0.748 Female 75.0 (7.5)

Table 3.3 – Site/Sex differences observed in morphometric measures and associated p-values resulting from two-sample Student’s T-test. All α = 0.05.

109

Observed Expected Location Sex Chi-Square p-Value Frequency Frequency

Male 37 42.5 Site 1 1.424 0.233 Female 48 42.5

Male 50 51 Site 2 0.039 0.843 Female 52 51

Table 3.4 – Observed gender differences for E. complanata and resulting statistics following Chi-

Square test (α = 0.05).

110

Figure 3.4 – Observed gender ratios as determined by biopsy analysis and expected counts for E. complanata at Sites 1 and 2 (nsite 1 = 85; nsite 2 = 102).

111

Site 1 Site 2 100

e 80

b 60

e 40

P 20

0 Stage g e g e e g g e g e e g n r n v r n n r n v r n i u i ti u i i u i ti u i p at b c at b p at b c at b o r A r o r A r el M so M so el M so M so v e te e v e te e e R a R e R a R D t/ L t/ D t/ L t/ en en en en Sp Sp Sp Sp

Sex le le le le a a a a m M m M Fe Fe

Percent is calculated within levels of sex.

Figure 3.5 – Gametogenic stages observed for E. complanata of both sexes at the two Cacapon

River study sites in May 2013. For each site, n = 5/sex.

112

Site 1 Site 2 100

e 80

b 60

e 40

P 20

0 Stage g al al g g al al g in ic ic in in ic ic in b p p b b p p b r y y r r y y r so t t so so t t so e A A e e A A e R y e R R y e R / rl t / / rl t / t a a t t a a t en E L en en E L en Sp Sp Sp Sp

Sex le le le le a a a a m M m M Fe Fe

Percent is calculated within levels of sex.

Figure 3.6 - Gametogenic stages observed for E. complanata of both sexes at the two Cacapon

River study sites in late July/early August 2013. Site 1 nmale = 6, nfemale = 3. Site 2 nmale = 6, nfemale =

113

Spring Summer 100

F 60

A 40

Site Site 1 Site 2 Site 1 Site 2 Site 1 Site 2 Site 1 Site 2 Sex Female Male Female Male

Panel variable: Season

Figure 3.7 – Average AAF values calculated for E. complanata at each site included in this study

(Males nspring = 10, n summer = 12; Females nspring = 10, nsummer = 7)

114

Spring Summer

A 40

Site Site 1 Site 2 Site 1 Site 2 Site 1 Site 2 Site 1 Site 2 Sex Female Male Female Male

Panel variable: Season

Figure 3.8 – Average GAF values calculated for E. complanata at each site included in this study

(Males nspring = 10, n summer = 11; Females nspring = 10, nsummer = 7). One outlier value was omitted for

Males, Site 1 from the Summer sampling.

115

)

( 2

s 1

M 0

-1 R1 Exp2 Exp1 R2 Group

Individual standard deviations are used to calculate the intervals.

Figure 3.9 – Average 20-month mass changes (g) for control (R1 and R2) and relocated (Exp1 and

Exp2) E. complanata (nR1 = 16, nR2 = 14, nExp1 = 11, nExp2 = 13) in the Site 1/Site 2 relocation study.

116

Figure 3.10 – Tukey pairwise comparisons for differences of means (g) for 20-month mass changes in E. complanata relocated between Site 1 and Site 2 (nR1 = 16, nR2 = 14, nExp1 = 11, nExp2

= 13).

117

3.5

3.0

s b 2.5

C 2.0

1.5 -11.311 -11.010 -10.709 -10.408 -10.107 -9.806 -9.505 -9.204 -8.903 -8.602 -8.301 -8 log(E2 Standard Concentration (M))

Individual standard deviations are used to calculate the intervals.

Figure 3.11 – 17β-estradiol standard (positive control) response data generated from POCIS extract analyses. Each point represents E2 standard response data from each of six test plates.

118

POCIS Sample E2 EC50 (M) E2 EC50 95% CI E2 EC20 (M) E2 EC20 95% CI

Site 1 Oct 2011 3.54E-09 2.69E-09 – 4.65E-09 1.79E-09 1.51E-09 – 2.12E-09

Site 2 Oct 2011 2.27E-09 1.30E-09 – 3.98E-09 1.01E-09 6.94E-10 – 1.48E-09

Site 1 June 2012 7.88E-10 7.05E-10 – 8.81E-10 3.71E-10 3.37E-10 – 4.08E-10

Site 2 June 2012 8.73E-10 6.88E-10 – 1.11E-09 4.23E-10 3.61E-10 – 4.95E-10

Site 1 Oct 2013 7.25E-09 1.07E-09 – 4.91E-08 2.20E-09 1.17E-09 – 4.14E-09

Site 2 Oct 2013 2.75E-09 1.83E-09 – 4.13E-09 1.46E-09 1.09E-09 – 1.96E-09

Table 3.5 – EC50 and EC20 values and associated 95% confidence intervals for 17β-estradiol standard dilution series curves from each POCIS extract YES assay microplate.

119

POCIS Sample Max Absorbance E2 Equivalent Concentration (M)

Site 1 Oct 2011 2.36 1.63E-09

Site 2 Oct 2011 1.97 4.21E-10*

Site 1 June 2012 1.80 3.12E-10

Site 2 June 2012 1.50 < 7.33E-12**

Site 1 Oct 2013 1.98 5.85E-10*

Site 2 Oct 2013 1.96 8.43E-10*

Table 3.6 – Maximal absorbance values observed for each POCIS composite extract and corresponding E2 Concentrations (M) from standard dilution series analyses. *Non-sigmoidal responses suggestive of toxicity at upper extract concentrations may influence max absorbance observed. **LOD listed when maximal absorbance failed to surpass upper 99% CI for negative control.

120

2.0000E-09

)

( 1.5000E-09

n 1.0000E-09

2 5.0000E-10

0.0000E+00

1 1 1 1 12 12 13 13 0 0 0 0 0 0 2 2 2 2 2 2 ct ct e e t t n n c c O O u u O O 1 2 J J 1 e 1 2 2 t te te e i i te e i it S S i it S S S S

Individual standard deviations are used to calculate the intervals.

Figure 3.12 - E2 concentrations (M) corresponding to average maximal responses (corrected absorbance) of POCIS extracts. Bars indicate 95% confidence intervals. Site 2 June 2012 data point represents upper limit of 99% CI for negative control, as absorbance values were below LOD.

121

Image 3.6 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post initial exposure to 17β-estradiol at the lowest concentration tested (1.9 x 10-5 ng/µL) showing viable ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n =

4) at this level of exposure.

122

Image 3.7 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post initial exposure to 17β-estradiol at the intermediate concentration tested (1.9 x 10-2 ng/µL) showing viable and nonviable ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n = 4) at this level of exposure.

123

Image 3.8 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post initial exposure to control solvent (27% EtOH) showing viable and nonviable ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n = 5) serving as controls.

124

Image 3.9 – Unstained portion of female E. insulsa biopsy tissue collected 10 days post initial exposure to 17β-estradiol at the highest concentration tested (19 ng/µL) showing atretic/resorbing ova. No viable ova were observed for three of five individuals at this level of exposure.

125

35000

30000

) 2 25000

(

a 20000

a v 15000

- 10000

5000

0 Control Low Medium High Exposure Group

Figure 3.13 – Median, Inter-quartile range, and total range of observed ova areas (µm2) from biopsy samples collected at time = 10 days following initial dosing of organisms (n = 5/group,

Control and High dose; n = 4/group Low and Intermediate dose) with E2.

126

Groups Compared Mean AAF Difference 95% CI Adjusted p-Value

Low Dose – Control -1.51 (-8.42, 5.40) 0.921

Medium Dose – Control 1.86 (-5.05, 8.77) 0.864

High Dose - Control 4.95 (-1.56, 11.47) 0.180

Table 3.7 – Mean differences in female AAF values observed for acini within organisms exposed to the three levels of 17β-estradiol versus solvent control (n = 50 acini included in the analysis for

Control and High Dose; n = 40 acini included in the analysis for Medium and Low Dose).

127

70.0

67.5

65.0

F 62.5

60.0

57.5

55.0

CTRL Low Mid High Exposure Group

Individual standard deviations are used to calculate the intervals.

Figure 3.14 – Average AAF values observed for E2 exposed E. insulsa versus controls. Bars represent 95% confidence intervals (n = 50 acini for control and High dose groups; n = 40 acini for

Low and Intermediate dose groups).

128

Image 3.10 – Stained portion of female E. insulsa biopsy tissue collected 5 months post initial exposure to control solvent (27% EtOH) showing viable mature and developing ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n = 5) serving as controls.

129

Image 3.11 – Stained portion of female E. insulsa biopsy tissue collected 5 months post initial exposure to 17β-estradiol at the lowest concentration tested (1.9 x 10-5 ng/µL) showing developing (D), mature (M), and atretic (A) ova and other cellular debris collected in the biopsy.

Viable ova were observed for all individuals (n = 4) at this level of exposure. Observed ova were classified according to appearance/staining characteristics, as well as areas measured and compared to threshold values previously identified for atretic, developing, and mature ova.

130

Image 3.12 – Stained portion of female E. insulsa biopsy tissue collected 5 months post initial exposure to 17β-estradiol at the intermediate concentration tested (1.9 x 10-2 ng/µL) showing developing, mature and atretic ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n = 4) at this level of exposure.

131

Image 3.13 – Stained portion of female E. insulsa biopsy tissue collected 5 months post initial exposure to 17β-estradiol at the highest concentration tested (19 ng/µL) showing developing, mature, and atretic ova and other cellular debris collected in the biopsy. Viable ova were observed for all individuals (n = 5) at this level of exposure.

132

50000

40000

)

(

a 30000

O 20000

5 10000

0 Control Low Medium High Exposure Groups

Figure 3.15 – Median, Inter-quartile range, and total range of observed ova areas (µm2) from biopsy samples collected at time = 5 months following initial dosing of organisms (n = 5/group,

Control and High dose; n = 4/group Low and Intermediate dose) with E2.

133

Figure 3.16 – 95% confidence intervals for the difference of E2 exposure level mean and control mean % atretic values (Dunnett Multiple Comparisons with a Control test) for female E. insulsa.

134

Biopsy Histology 60

v r 40

t 30

P 20

0 Late Active Late Atypical Mature Late Active Late Atypical Mature Gametogenic Stage

Panel variable: Histo Biopsy Appraiser Percent is calculated within all data.

Figure 3.17 – Gametogenic stage determinations for male E. insulsa sampled 5-months post initiation of E2 exposure study (n = 20) using the biopsy-based approach as well as the histological approach (see Figures 2.3 and 2.8 for methodologies).

135

Figure 3.18 – Dunnett Simultaneous 95% CIs. Intervals represent 95% confidence intervals for the difference between 5-month post-exposure mean AAF values among Control organisms (n = 5) and mean AAF values among organisms at each exposure level (n = 5 organisms per exposure level).

136

3.0

2.5

i 2.0

r t 1.5

a 1.0

0.5

0.0 CTRL High Intermediate Low Exposure Level

Figure 3.19 – Boxplots of average proportion (%) of atretic tissue observed within measured acini in E. insulsa (n = 5/group) 5-months post E2 exposure. Significance (α= 0.05) was observed for Intermediate (p = 0.029) and Low (p = 0.015) level exposure groups.

137

)

o 20

(

s 15

r 10

% 5

Control High Intermediate Low Exposure Group

Figure 3.20 - Boxplots of average proportion (%) of atretic tissue observed in biopsy samples analyzed from E. insulsa (n = 5/group) 5-months post E2 exposure. Significance (α= 0.05) was not observed for any group (p = 0.261).

138

Chapter 4: Investigation into recent observations of recruitment failure in the East Fork West

Branch St. Joseph River population of the federally endangered freshwater mussel, Pleurobema

clava (Bivalvia: Unionidae)

Introduction

Population declines in a variety of aquatic species have been well documented over the last century, and there is no better example of this than among those species within the order

Unionoida. North America is home to the richest assemblage of unionids in the world, however an alarming number of the nearly 300 native species are, to some degree, imperiled (TNNMCC,

1998). Within the last 100 years, 29 species, or 10% of the unionid taxa found in the United States

(US), have become extinct (Haag and Williams, 2014). As of April, 2015, 76 US species have been listed as federally endangered and another 13 are considered threatened (USFWS, 2015). This amounts to a federally listed status for approximately 30% of all US species. As independent assessments have expressed concern for 199 of the nearly 300 US species (Haag and Williams,

2014), and recent estimates have suggested that unionid species may soon experience an extinction rate of up to six percent per decade (Ricciardi, 1999), there is an urgent and obvious demand for focused research and conservation efforts within this group.

Thought to have once been common and widely distributed throughout the Ohio River and Lake Erie drainages, the clubshell mussel, Pleurobema clava (Lamarck, 1819), received federal protection under the Endangered Species Act in February 1993, and is now believed to be limited to only a few (approximately 8 to 10), generally small, isolated populations in Ohio, Michigan,

139

Indiana, Pennsylvania, and West Virginia (USFWS 1994). One of these populations exists in the

East Fork West Branch St. Joseph River in southwestern Michigan. The most recent observational data collected on this population indicated a complete lack of recruitment, and described this population as relict, consisting of only a few large, mature individuals (G. Thomas Watters, personal communication). This situation is not unique, as a hallmark of many of the enigmatic population declines that have occurred since the 1970s has been an apparent lack of recruitment, leaving behind small populations consisting only of a few aging individuals (Strayer and Malcolm,

2012; Haag and Williams, 2014).

Due to the complex nature of the unionid life cycle, several possibilities exist at which reproductive failure within a population may occur. While detailed descriptions and discussions of unionid biology and reproduction may be found elsewhere (Watters, 2007), briefly, it is generally accepted that two differing reproductive strategies exist within the unionids, the differentiating factor between these approaches being the duration of glochidial brooding. The general life cycle of all unionids, while complex, appears to be, essentially, the same. While a few hermaphroditic and “occasionally hermaphroditic” species have been observed (Van Der Schalie,

1966; Heard, 1979; Kat, 1983), unionid species are typically dioecious, with adults developing either ova or sperm in diffuse gonadal tissues located throughout the visceral mass. Though males release sperm freely into the water column, often packaged by the thousands into specialized structures referred to as, “sperm balls” or spermatozeugmata, fertilization occurs internally.

Females siphon male gametes from the water and fertilization of ova takes place within the suprabranchial chamber of the gill. Fertilized ova are then moved to the gill marsupia and develop into an obligate parasitic life stage known as a glochidium, which requires an encystment period on a host organism, usually a fish, prior to transformation to the juvenile stage. Following

140 successful encystment and transformation on the appropriate host, the juvenile mussel drops to the substrate and continues growth and development to maturity.

Based upon the complex approach to reproduction employed, a variety of points exist at which reproductive failure in unionids could occur. Regardless of the point of failure, several potentially causative factors have been proposed for these situations in which unionid populations are in apparent decline, yet no obvious cause can be identified. This short list of hypothesized contributors includes sedimentation, disease, and exposure to anthropogenic chemicals such as pesticides and ammonia (Haag and Williams, 2014), in addition to presence and quality of suitable hosts for glochidial encystment (Watters, 2007). In regards to the exposure hypothesis, a growing body of evidence is beginning to demonstrate a possible inordinate sensitivity of mussels, particularly during early life stages, to these compounds (Bringolf et al.,

2007a; Bringolf et al., 2007b; Bringolf et al., 2007c; Newton and Bartsch, 2007; Wang et al., 2008;

Strayer and Malcom, 2012). Less is known, however, about possible effects of contaminant exposure on gametogenesis and gamete viability in adults, and whether this could act directly, or possibly contribute some additive effect, to situations in which recruitment appears to have ceased.

The East Fork West Branch St. Joseph River is located in the headwaters region of the St.

Joseph River Watershed in Hillsdale County, Michigan. This watershed encompasses 694,400 acres throughout Michigan, Ohio, and Indiana (SJRWI, 2012). Land use in the watershed is primarily agricultural, as nearly 80% of the watershed’s acreage is dedicated to cropland or pasture, and previous investigations into water quality throughout the watershed have indicated periodic spikes in surface water concentrations of each of four common-use herbicides sampled, which seemed to align with spring herbicide applications and heavy-rain events in the area (2012).

141

This same pattern of intermittent contaminant influx may occur for other commonly employed pesticides, many of which are known EDCs and xenoestrogens, though no recent or additional data have been collected to support or refute this hypothesis.

The aim of this study is to verify the previously observed lack of recruitment in the East

Fork West Branch St. Joseph River P. clava population and to identify the stage at which the complex reproductive strategy of these organisms is failing. Additionally, as detectable levels of herbicide residues have been found in surface waters throughout the Maumee River Basin, including the East Fork West Branch St. Joseph River (2012), seasonal measures of total surface water estrogenic activity are utilized in order to build a weight-of-evidence argument for the cause(s) and level(s) of disruption that have contributed to failed reproduction in this P. clava population.

Materials and Methods

Site Selection and Field Sampling

Two study sites were included in this study. The primary study site (EF) is located in the

East Fork West Branch St. Joseph River, at approximately 41° 46’ 53.3” N, 84° 39’ 02.5” W

(elevation 306.3 m) along Woodbridge Road, approximately nine miles south of the town of

Hillsdale, Hillsdale County, Michigan (MI). This site had been previously identified as a location in which P. clava were present, but recruitment had ceased. The second site (AL), selected as a

“reference” for comparison, was located at approximately 41°28’22.1” N, 79°29’56.5” (elevation

314.9 m), along the right descending bank of the Allegheny River, directly beneath the Route 62 crossing in Tionesta, Forest County, Pennsylvania (PA). This site was selected as a “reference” in this study, as it represents one of the only remaining viable, reproducing populations of P. clava known to exist. Due to the federally endangered status of this species, care was taken to minimize

142 disruption of substrate during sampling, and limited numbers of organisms were collected via hand-grab and visual identification using glass-bottom buckets. A second, surrogate species, E. dilatata, was sampled concurrently. This species was selected to serve as a surrogate as it represented the only common short-term glochidial brooding, or tachytictic, unionid species available for significant collection at both sites. Organism sampling occurred between May 2012 and October 2013. During each sampling event, water temperature and pH were recorded and P. clava were collected and held in-stream for nonlethal sampling of the developing gonad using the biopsy technique described earlier (see Methods – Chapter 2), though a pre-loaded 20-gauge syringe was used in place of the typical 18-gauge syringe in order to minimize risk of injury to organisms sampled. Individuals observed were marked (shell etch) weighed (g), and maximum valve length (mm) was recorded prior to biopsy collection. Biopsy samples were collected only once for any given individual. A similar approach was taken for sampling of E. dilatata. Organisms at AL were found along the right descending bank in a shallow run exhibiting a high density of P. clava, while E. dilatata generally required more sampling effort to collect in sufficient numbers.

P. clava were observed more sporadically at EF, but were generally located in shallow runs, while

E. dilatata were common. Gametogenic stage was determined for individuals biopsied during

2013 sampling events according to procedures described in Figure 2.8 (Males) and Table 2.10

(Females). Due to limited sample sizes, staged individuals were correspondingly classified as either “viable” or “non-viable” according to whether active gametogenesis was or was not observed, and these classifications were employed for site comparisons by Fisher’s Exact test.

Sampled organisms were also evaluated for evidence of gravidity by visual inspection of gills, as the marsupium of gravid females generally appeared enlarged or swollen. Up to three gravid females at each site were selected for collection of glochidia for viability testing. Glochidia were flushed from water tubes using a water-filled 18-gauge syringe and were collected in a petri

143 dish at the site. Collected glochidia from each individual were divided into three sub-samples and were observed under a dissecting microscope in order to initially determine degree of development, as well as to assess the proportion of individuals having open or closed valves.

Glochidial viability was then immediately tested for fully developed individuals using a procedure modified from Wang et al (2007) and suitable for the field. Briefly, glochidia were divided into three sub-samples and held in petri dishes with water flushed through the water tubes during collection. Individuals were initially observed on a dissecting microscope (10x) and the number of glochidia having closed valves was recorded. Several drops of a saturated NaCl solution were added to each petri dish and the number of organisms having closed valves was again recorded.

Viability (%) of each subsample was calculated as 100 X (nclosed+NaCl – nclosed-NaCl) / ntotal glochidia), where nclosed+NaCl is the number of individuals with closed valves after NaCl exposure and nclosed-NaCl is the number of individuals with closed valves before exposure to the NaCl solution (2007).

The presence or absence of suitable host species for the glochidia of P. clava was investigated by manually seining at each site in late June 2013. Netted fish known or suspected to be potential P. clava hosts were visually inspected for evidence of glochidial encystment on fins immediately upon collection and most were subsequently released. Up to three individuals of each known or suspected host species identified at each site were taken, transported to the laboratory on ice, and preserved in 95% EtOH for microscopic inspection of fins and gills in order to more accurately investigate possible evidence of successful glochidial encystment.

Site differences in gametogenic stage, sex ratio, and organism size (total organism wet weight and maximum valve length), were determined when comparative data between sites were available, by applying appropriate transformations, when required, and using the two-sample

144

Student’s T-test when all parametric assumptions, including normality, were met. Otherwise, comparisons were made using Mood’s Median, Chi Square, or Fisher’s Exact tests.

POCIS Sampling and Yeast Estrogen Screen Extract Analysis

AQUASENSE-P Polar Organic Chemical Integrative Sampling Devices (POCIS) (EST

Laboratories, St. Joseph, MO), described earlier, were utilized in this study, and attempts were made to deploy samplers seasonally for 60-day sampling periods at each study site. Realized deployment periods are listed in Table 4.1. Oasis HLB AQUASENSE-P samplers were utilized for all deployments in accordance with guidelines published for environmental monitoring studies

(Alvarez, 2010) and all deployments, sample extractions, and analyses were carried out according to methods and procedures described earlier (see Methods – Ch. 2 – Passive Surface Water

Sampling and Yeast Estrogen Screen) unless otherwise stated. A field blank POCIS disc (n = 1 per sampling event per site) was employed for each sampling deployment/retrieval and utilized for quality control purposes in order to ensure that deployment, retrieval, storage, processing, and analysis did not have a significant effect on deployed discs. Blank discs were returned to their original containers and stored frozen during the deployment period.

Pooled (3-disc) concentrated POCIS extracts were analyzed for total estrogenic potency with the recombinant yeast estrogen screen (YES) previously described. Samples of either 10 µL

POCIS extract + 190 µL solvent (2011 and 2012 samples) or 200 µL POCIS extract (2013 samples) were added to the first column of a 96-well microplate in triplicate, serially diluted across the plate using pure solvent (MeOH), and allowed to evaporate to dryness prior to addition of assay media and inoculation of plate wells with freshly cultured recombinant yeast. An E2 standard positive control was employed in the first row of each test plate and negative controls were

145 included between the positive control and all sample rows to minimize any potential cross- contamination of sample wells. Upon visual observation of adequate response (color development) in the positive control following incubation, each plate was unsealed and absorbance in each well was measured at 540 nm (colorimetric response) and 620 nm (turbidity) on a BMG Fluostar OPTIMA microplate reader. Data collected were analyzed for estrogenic response and EEQ values, when appropriate, were determined using Microsoft Excel and

GraphPad Prism analytical software. In instances where EEQ values could not be determined using

Equation 1 (see Results section, Chapter 3), response data were used to determine an observed maximum E2 standard equivalent concentration for the sample. The two-sample Student’s T-test was used for site extract estrogenicity comparisons when all parametric assumptions, including normality, were met. Otherwise, comparisons were made using Mood’s Median test.

Results

Morphological Characteristics

Total organism mass (g) and maximum valve length (mm) were recorded for a total of 30

P. clava at EF and 42 P. clava at AL. Distributions for each measure are presented in Figures 4.1 and 4.2. Mass of EF P. clava (Image 4.1) ranged from 2.0 g – 96.4 g (Median = 45.9 g), while maximum valve length ranged from 20.41 mm – 79.77 mm (Median = 56.06 mm). Similarly, mass of AL P. clava ranged from 2.0 g – 98.3 g (Median = 36.4 g), while maximum valve length ranged from 19.15 mm – 82.81 mm (Median = 55.95 mm). No differences in P. clava mass (Mood’s

Median Test: p = 0.633, α = 0.05) or length (Mood’s Median Test: p = 1.000, α = 0.05) were detected between sites. The distributions of both mass and valve length at EF, however, appeared somewhat bimodal (Figure 4.1), while those for organisms sampled at AL appeared to be more

146 representative of normal curves (Figure 4.2). E. dilatata populations sampled concurrently with

P. clava at each site demonstrated normal distributions for both mass (g) and maximum valve length (mm) at each site (Figures 4.3 and 4.4). E. dilatata mass at EF ranged from 26.8 g to 96.2 g

( x ̅ = 57.7 g; SD = 20.4 g), while valve length ranged from 55.36 mm to 96.10 mm (x ̅ =78.00 mm;

SD = 9.86 mm). At AL, mass ranged from 7.6 g to 44.5 g (x ̅ = 28.6 g; SD = 8.34 g), while valve length ranged from 44.13 mm to 80.28 mm (x ̅ = 62.74 mm; SD = 9.48 mm). These differences in mass and maximum valve length observed between EF and AL E. dilatata were significant (two sample

Student’s t-test: pmass <0.001, α = 0.05; plength <0.001, α = 0.05).

Seasonal Gametogenesis

Gametogenic assessments were conducted on biopsy samples collected between May 2013 and

October 2013. Of the P. clava sampled in May 2013, six males and two females were observed at

EF, while two males and nine females were observed at AL. Four new organisms were sampled from EF in July 2013 (two male, two unidentified), followed by an additional four individuals (three male, one female) in late September. At AL, three males and six females were observed in July biopsy samples, while four males and six females were found in the early October sampling.

Seasonal gametogenic stages observed for both male and female individuals at EF and AL are shown in Figures 4.5 and 4.6, respectively. EF males exhibited a wide range of developmental stages in May, including Inactive, Early Active, Late Active, Early Atypical, and Spent/Resorbing, while only Late Active development was observed for AL males at that time. Subsequent EF sampling events for P. clava saw one Early Atypical and one Spent/Resorbing male in July, followed by two Early Atypical males and one Late Atypical male in late September. At AL, July and early

October sampling of P. clava resulted in classifications of one Late Active and two Mature

147 organisms (July), while all males observed in Autumn (n = 4) were considered Early Atypical. The two female P. clava sampled from EF in May were classified as Developing and Spent/Resorbing, while those at AL exhibited characteristics of Developing, Mature, and Spent/Resorbing gametogenic stages. Summer sampling resulted in the collection of no females at EF, though the gender of two individuals classified as Inactive was not positively identified, as these biopsy samples contained no identifiable reproductive cells. Gametogenic stages of six females collected from AL in July were split evenly between Mature and Spent/Resorbing (n = 3 per stage). Only

Spent/Resorbing females were observed in samples from both sites in late September/early

October. Due to low sample sizes and the, relatively, high number of potential gametogenic classifications possible, the stage identified for each individual was further designated as either

“Viable” or “Non-viable” and male and female samples from each site were pooled (nEF = 7, nAL =

10). The “Non-viable” designation was assigned to Inactive and Spent/Resorbing individuals, as these samples exhibited no indication of the active production of viable reproductive cells at the time. All other stage classifications were considered “Viable” (nEF = 9, nAL = 20). These data for P. clava were compared for each sampling season of 2013 (Table 4.2), and no significant differences were detected between sites for any sampling period (Fisher’s Exact Test pspring = 0.603; psummer =

0.217; pautumn = 0.559, all use α = 0.05). Similar results were obtained following gametogenic staging, viability determinations, and analysis of E. dilatata biopsy data (Table 4.3). A total of 18 individuals of this species were considered “Non-viable” at EF, while 22 were found to contain

“Viable” gametes. E. dilatata samples at AL consisted of 6 “Non-viable” and 22 “Viable” organisms. E. dilatata between-site comparisons were made for each sampling season, and no significant differences in gonad viability determinations were detected (Fisher’s Exact Test pspring

= 0.685; psummer = 266; pautumn = 0.350, all use α = 0.05). Gametogenic stages observed for E. dilatata and used to make viability designations are provided in Figures 4.7 (EF) and 4.8 (AL).

148

Sex Ratios

The sex of a total of 21 P. clava was positively identified between May 2012 and October

2013 at EF, while 36 individuals were sexed at AL during that period. No significant departure from an expected male-to-female ratio of 1:1 was observed for EF (Chi Square test: p = 0.827, α = 0.05), while this ratio differed significantly from 1:1 at AL (Figures 4.9, 4.10), as a total of 25 females and only 11 males were identified at that site (Chi Square test: p = 0.020, α = 0.05). Sex ratios were also calculated for E. dilatata at each site (Figures 4.11 and 4.12). Individuals collected and sexed at EF (n = 29) exhibited no significant departure from 1:1 (Chi Square test: p = 0.194, α = 0.05), nor did E. dilatata sampled at AL (Chi Square test: p = 0.414, α = 0.05; n = 24 organisms identified).

POCIS Deployments and Yeast Estrogen Screen Analysis of Extracts

POCIS sampling devices were deployed at each site seasonally between October 2011 and

August 2013 (see Table 4.1). Concentrated three-disc composite sampler extracts were assayed using the YES procedure described earlier. Sample extracts collected from 2011/2012 deployments generally exhibited the expected dose-dependent sigmoidal response (10 µL extract assayed), though all observed responses were weak in these samples and EC20 values were utilized for calculation of EEQ values (ng E2/POCIS). EEQ values for EF deployed samplers ranged from ND (no response above negative controls) to 0.44 ng E2/POCIS, while AL responses ranged from ND to 0.30 ng E2/POCIS. Higher extract volumes (200 µL) were assayed for samplers deployed during the 2013 sampling season due to the weak responses observed for 2012 assays, however, this extract volume resulted in a non-sigmoidal response curve in all extracts exhibiting a positive estrogenic response, and EEQ values were, therefore, not determined for these samples. A maximum absorbance E2 equivalent concentration was determined for each of these

149 samples in place of a calculated EEQ. EEQ values and maximum absorbance E2 equivalent concentrations in absence of EEQ data for deployed samplers are presented in Table 4.4.

Extracts of POCIS deployed at EF exhibited increasing estrogenicity response from winter

2011 (no detection) to spring 2012 (0.261 ng E2/POCIS EEQ) to summer 2012 (0.438 ng E2/POCIS

EEQ). Responses for AL POCIS extracts also exhibited the lowest EEQ during the winter 2011 deployment (0.040 ng E2/POCIS EEQ), however the spring 2012 extract exhibited a slightly increased EEQ value (0.304 ng E2/POCIS) in comparison with the summer 2012 EEQ response

(0.260 ng E2/POCIS). However, it is critical to note that the summer-deployed AL sampler was retrieved partially exposed, and the total submerged time is not known. The maximum E2 standard equivalent responses observed for 2013 extracts exhibited similar trends for each site among 2012 samples (Table 4.4). The highest observed E2 equivalent concentration for 2013 EF samples was found for the May – July deployment period (3.881 nM equivalent), while the highest recorded maximum equivalent E2 concentration observed for all samples was found for the AL

July – September 2013 deployment (7.202 nM equivalent).

Gravidity and Glochidial Viability

All P. clava and E. dilatata collected for biopsy during the study were visually inspected for evidence of gravidity. No gravid P. clava were observed at EF during any sampling event. One single female E. dilatata sampled in late July, 2013 at EF appeared gravid, though glochidia were not collected from this organism to verify the observation. At AL, visual inspection of the gill resulted in a total of three individuals (here, designated AL-1, AL-2, and AL-3) being suspected of gravidity in early July 2012, and an additional five suspected in late July, 2013. No E. dilatata sampled at AL were suspected of gravidity. Glochidia were flushed from the water tubes of the

150 three individuals observed during July 2012 sampling, in order to verify the observations and test viability of any glochidia collected. Mature glochidia were collected from two of these three individuals (AL-1 and AL-2), while all glochidia flushed from the third organism (AL-3) appeared immature and were not tested. These larvae exhibited a bell-like shape and appeared, as has been previously described for immature glochidia (Ingersoll et al., 2007) to remain fully encapsulated within the original egg membrane. The average observed incidence rate of mature glochidia having closed valves prior to addition of NaCl solution was low (AL-1 = 8.7 %, AL-2 = 2.1 %). Viability test results for mature glochidia collected from each gravid P. clava are presented in Table 4.5.

Host Presence and Glochidial Encystment

Efforts at each site were made to verify presence of, or alternatively support possible absence of, the blackside darter (Percina maculate), the striped shiner (Luxilus chrysocephalus), the central stoneroller (Campostoma anomalum) and the northern logperch (Percina caprodes), all of which have been experimentally identified as potential fish hosts for P. clava (O’Dee and

Watters, 1998). Organisms collected by seining/netting were also briefly inspected visually for any evidence of glochidial encystment. Seining events were conducted in early July, 2012 as well as late June, 2013. Observations of each known host species were made at both EF and AL, indicting suitable host presence at each site. No evidence of glochidial encystment was found for any organisms visually inspected in the field, nor for organisms of each species (n = 3/species/site) collected during the 2012 sampling event, for which gill and fin tissue were more adequately inspected using a dissecting microscope (10x).

151

Discussion

This study was undertaken following evidence of recent recruitment failure and cessation of successful reproduction in the East Fork West Branch St. Joseph River population of the federally endangered clubshell, P. clava. While reports of this population date back nearly a century (Winslow, 1918), later surveys have suggested that it is relatively confined to an approximately 10 mile stretch of stream in Hillsdale County, Michigan, and is characterized by large, aging individuals and an apparent lack of reproduction, evidenced by the absence of young adult, juvenile, or newly transformed organisms (Watters, 1988; G. Thomas Watters personal communication). If accurate, the nature and structure of this population places it at a high risk of future loss. The drainage in which this population exists has, however, been identified as one of

10 drainage basins in which viable P. clava populations must be established or re-established prior to any consideration of federal reclassification of this species (USFWS, 1994). Therefore, the primary objective of this study to identify the stage(s) at which reproductive failure was occurring among individuals comprising this population and, secondarily, to investigate the potential presence of xenobiotics in surface waters proximate to the relatively confined range of this endangered species in the East Fork West Branch St Joseph River that could pose a risk to past or present reproductive success.

Due to the complex nature of the unionid life cycle (see Watters, 2007), it was necessary to investigate several potential failure points in the P. clava reproductive pathway in order to attempt to ascertain the stage(s) at which reproduction was failing in this population. These include viable gamete production, fertilization, larval development and viability, and host presence, as these organisms utilize a parasitic larval stage requiring, typically, a fish host. Limited sampling was conducted in this population between spring 2012 and autumn 2013. Mass and

152 maximum valve length distributions for this population (see Figure 4.1) suggest that successful reproduction has occurred in very recent years. Distributions also may suggest, however, the possibility that a period of reduced reproductive success, or even complete reproductive failure, has been experienced for a time in this population. An apparent gap in mass and length distributions was observed among organisms sampled, in which organisms between approximately 50 mm and 60 mm in length and between approximately 20 g and 40 g in total weight were scarce and, nearly, absent altogether. This resulted in the production of what appears as a bimodal size distribution for this sample of P. clava in the East Fork West Branch St.

Joseph River. A similar distribution was not observed for E. dilatata sampled concurrently at the site. Additionally, while a similar gap appeared in size data for P. clava in the Allegheny River, this gap was at the highest end of the distribution, which, otherwise, appeared normal.

Unfortunately, the distribution cannot be verified as being fully representative of this P. clava population in the East Fork, as quantitative sampling procedures, such as those discussed in detail for unionid populations (see Strayer and Smith, 2003) were not permitted due to the listed status of this species. It was clear, though – based upon the observation of organisms ranging from approximately two to 100 grams in total weight and sampling of 10 P. clava weighing less than 20 g and measuring less than 45 mm in maximum valve length – that recent P. clava recruitment has succeeded in this population, likely within the last 3 to 5 years. Whether consistent, annual recruitment has occurred for any length of time or continues to occur at this time, however, remains unclear.

Data collected in this study are some of the first known data collected on gametogenesis, gravidity, and sex ratio from natural populations of P. clava. Here, based upon findings for organisms sampled in what was considered the “reference” population in the Allegheny River, it appears that P. clava contain ripening and mature gametes from late spring to mid-summer,

153 followed by resorption of the gonad (female) or atypical spermatogenesis (male) in autumn. Due to the highly endangered status of P. clava, no organism sacrifice was conducted for this species, and, therefore, no species-specific threshold values for gametogenic staging were defined.

Instead, those values identified for E. complanata were used as the only available alternative.

Gravidity was observed in late June and early July, and both mature and developing glochidia were observed in separate individuals in early July 2012. The sex ratio observed for P. clava at AL exhibited significant departure from an expected 1:1 ratio and was dominated by females (M:F ratio = 0.44). This, again, could not be verified for the population, as haphazard organism sampling at the substrate surface was employed, and no quantitative sampling was conducted in this study.

In comparison, no mature females were observed in 2013 EF samples staged for gametogenesis, though, in total, only three females were observed at this site (two in May and one in September).

Male P. clava staged from EF, however, exhibited a high degree of variability in gamete development during Spring 2013 sampling, as of the six individuals assessed, stages assigned included Inactive, Early Active, Late Active, Early Atypical, and Spent/Resorbing, while no mature individuals were observed. Further sampling during July and September 2013 saw only atypical spermatogenesis (both Early Atypical and Late Atypical) and Spent/Resorbing stages represented among males at EF. Regarding E. dilatata sampled from EF in 2013 and staged, females exhibited a high rate of the Spent/Resorbing classification in each of the three sampling events, while males appeared to be successfully undergoing spermatogenesis and maturing in spring, while initiating atypical spermatogenesis in summer and continuing this process into autumn. As the population density of P. clava at EF appeared to be low, particularly in comparison to that observed (but not quantified) at AL, any degree of developmental asynchrony or impairment of gamete development in this population could be of significant consequence to successful recruitment, as studies of unionid species have implicated population density, alone, as playing a significant role

154 in reproductive success (Strayer et al., 2004). A lack of expected gametogenic development for P. clava sampled and staged at EF aligns with an absence of observed gravidity among females at that site, while development and maturation of gametes with rising water temperatures in spring at AL, as is generally expected for most unionid species, aligns with the observation of successful fertilization and gravidity among females at that site. However, while obviously important, a great number of other factors outside successful gamete development likely play a role in the rate of fertilization and glochidial development in any unionid population, such as population density, referenced above. Other known factors that could impact successful reproduction in P. clava include, but are in no way limited to, sedimentation, disease, and exposure to anthropogenic chemicals such as pesticides and ammonia (Haag and Williams, 2014). In addition, the reproductive success of these species are inextricably tied to the presence of suitable hosts for glochidial encystment and transformation, and any efforts in conservation or investigations into ecotoxicological effects in these organisms must also focus attention on the host(s) (Watters,

2007). In this study, host presence was evaluated at both sites, and, while no evidence of glochidial encystment was observed for fish collected, the presence of known P. clava hosts at each site was, at minimum, verified.

In order to assess the presence of xenobiotic compounds potentially capable of modulating or inhibiting gamete development and reproductive success in P. clava populations residing at each site in the study, POCIS samplers were deployed seasonally and extracts were assayed using the YES procedure in order to determine whether estrogenic compounds were present in surface waters. Estrogenic responses were low relative to the E2 standard concentration series employed for each test plate. However, when sample volume was increased from 10 µL assayed to 200 µL assayed following initial sample responses just above negative controls, an unexpected result was apparent toxicity to yeast or inhibition of colorimetric

155 response at the highest concentrations in the dilution series. This produced non-sigmoidal response curves and EEQ values were not calculated for these (late 2012 and 2013) sample extracts, though presence/absence of estrogenic compounds in the sampler extracts was still assessed.

Extracts from EF deployed samplers exhibited some level of estrogenicity above negative controls in all extracts except the first, deployed from October 2011 through December 2011.

Sampler extracts from AL deployed POCIS indicated presence of estrogenic compounds in the

Allegheny River in all extracts except that for the POCIS deployed in October 2012. This sampler, however, due to water levels and flow conditions, was unable to be retrieved until the following sampling event in May 2013. Passive samplers, such as the POCIS, are designed for long-term monitoring and deployment, and deployment periods of up to one year have been performed

(Alvarez, 2010). However, deployments are generally limited to two to three months as issues with biofouling, vandalism, and changes in sampling kinetics over time are possible (2010).

Whether any such issues effected the overall estrogenic potential of the samplers deployed for the period between October 2012 and May 2013 was not investigated, though no significant biofilm or biofouling was observed on collected sampling discs or deployment canisters.

Additional complications were experienced for both the EF and AL samplers deployed in

2012. The EF sampler deployed in August 2012 was likely stolen or dislodged from the site and carried downstream by a high-flow event, and could not be found. The AL sampler deployed in

June 2012 was recovered, but was found partially exposed at the time of retrieval, and the actual fully submerged exposure time of this sampler is unknown beyond the last verified observation of the undisturbed sampler on day 22 of the deployment.

156

No additional testing of sampler extracts was conducted to determine what compound(s) may have been responsible for the estrogenic responses observed. The area in which this stream is located is typically associated with agricultural land use activity, and reports of pesticide contamination have been published for waters in the region, including the East Fork West Branch

St. Joseph River (SJRWI, 2012). Noted chemicals evaluated include alachlor and metalochlor, as well as atrazine. It is important to note that total estrogenicity was assayed for POCIS extracts in the present study, and many chemicals used commonly in agricultural practice exhibit some degree of estrogenicity, including alachlor (Klotz et al., 1996). However, other compounds often used in agriculture with the potential for surface water contamination have no estrogenic interaction, or may even exhibit anti-estrogenic activity, as has been reported for atrazine, though this activity has not been observed in all studies testing its potential effects (Eldridge et al., 2008).

While these compounds have been reported to contaminate surface waters in this region, any remarks on their potential association with estrogenic responses observed for POCIS extracts in this study is purely speculative.

The EF site is located in the headwaters region of the St. Joseph River, while the AL site is located in the main stem of the Allegheny River. While temperature and pH data were recorded for the two sites in this study and were similar seasonally (data not shown), flow characteristics, substrate composition, sedimentation, and a variety of other factors exist that could possibly confound any perceived correlations between presence of dissolved estrogenic compounds in surface waters and impacted reproductive success of P. clava. Direct site comparisons for estrogenic response of POCIS extracts are only recommended when an estimate of the response’s equivalent water-based concentration of 17β-estradiol can be determined for each POCIS sampler. To calculate this estimated concentration, the sampling rate (Rs) value(s) for the chemical(s) of interest must be known (Alvarez, 2010). In the absence of these values, and, also,

157 because experimentally derived values can change based upon a variety of environmental factors, considering the observed differences between the two sites, no direct comparisons of EEQ or maximum observed E2 equivalent concentrations between sites were made. Even seasonal comparisons for a given site should be interpreted with caution due to the high degree of variability in flow conditions that are possible throughout the year. That said, this study demonstrated the presence of some chemical(s) in surface waters of both P. clava populations.

While the compounds responsible for this activity are unknown, regardless of the particular compound or compounds responsible for this activity, estrogenic compounds have demonstrated an ability to elicit physiological changes in gonad development and reproductive maturation of freshwater mussels and other freshwater invertebrates (Hardege et al., 1997; Gagné et al., 2001;

Regoli et al., 2001; Gagné and Blaise, 2003; Jobling et al., 2004; Nentwig, 2007; Ortiz-Zarragoitia and Cajaraville, 2006) and it is recommended that future conservation efforts for species such as

P. clava take the presence of these compounds into consideration in developing approaches to organism recovery.

It is apparent that earlier reports of a complete cessation of reproduction in the East Fork

West Branch St. Joseph River were either made in error or, more likely, associated with some temporary pause in reproduction within this population. Evidence of recent recruitment was observed in this study. However, also observed for P. clava in the East Fork population were alarming characteristics of gamete development suggesting high rates of resorption and highly variable development among individuals within the population. These observations may represent the earliest level at which reproduction is, or has been, impaired for P. clava in this stream. In addition, no evidence was found to suggest current reproductive success beyond gamete development in this population, as no gravid females were observed (lack of fertilization) and none of the known host fish collected exhibited any signs of recent glochidial encystment.

158

Based upon these findings, as well as those in support of the presence of xenobiotics potentially capable of influencing reproduction and gamete development in aquatic organisms, while this population may be experiencing occasional reproductive success during any given year, the long- term outlook for the viability of this population is still in question.

159

Sampler Deployment Date EF Deployment Period (days) AL Deployment Period (days)

October 2011 56 56

April 2012 65 65

June 2012 60 >22, <60*

August 2012 NA** 62

October 2012 219 222

May 2013 60 60

July 2013 60 60

Table 4.1 – Deployment periods for POCIS samplers at both EF and AL study sites. *Sampler at

AL was found partially exposed near the river bank following the June 2012 deployment. Actual time submerged is unknown beyond site visit on day 22. **Sampler at EF was not located following the August 2012 deployment.

160

Image 4.1 – Examples representing the size range observed for P. clava in the East Fork West

Branch St. Joseph River. This size range provides evidence of recent successful recruitment within this population.

161

e 6

r 5

v 4

s b 3

0 0 20 40 60 80 100 24 32 40 48 56 64 72 80 Mass (g) Length (mm)

Figure 4.1 – Total mass (g) and maximum valve length (mm) distributions for P. clava at EF. Each figure represents data for a total of 30 organisms sampled between Spring 2012 and October

2013.

162

O 5

0 0 20 40 60 80 100 20 30 40 50 60 70 80 Mass (g) Length (mm)

Figure 4.2 – Total mass (g) and maximum valve length (mm) distributions for P. clava at AL. Each figure represents data for a total of 42 organisms sampled between Spring 2012 and October

2013.

163

e 10

s 6

O 4

0 20 40 60 80 100 60 70 80 90 100 Mass (g) Length (mm)

Figure 4.3 - Total mass (g) and maximum valve length (mm) distributions for E. dilatata at EF.

Each figure represents data for a total of 20 (mass) and 32 (length) organisms sampled between

Spring 2012 and October 2013. Sample size discrepancies are due to equipment failure during

July 2013 sampling, which prevented collection of mass data for E. dilatata sampled that day.

164

y c 8

b 4

0 10 15 20 25 30 35 40 45 45 50 55 60 65 70 75 80 Mass (g) Length (mm)

Figure 4.4 - Total mass (g) and maximum valve length (mm) distributions for E. dilatata at AL. Each figure represents data for a total of 36 organisms sampled between Spring 2012 and October

2013.

165

Sampling Season Site n per sex Total n p-value

4 (M Viable) 2 (M Non- 5 Viable EF viable) 3 Non-viable 1 (F Viable) 1 (F Non-viable) Spring 2013 0.603 2 (M Viable) 0 (M Non- 9 Viable AL viable) 2 Non-Viable 7 (F Viable) 2 (F Non-viable) 1 (M Viable) 1 (M Non- viable) 1 Viable EF 0 (F Viable) 3 Non-viable 0 (F Non-viable) 0 (U Viable) Summer 2013 0.217 2 (U Non-viable) 3 (M Viable) 0 (M Non- 7 Viable AL viable) 2 Non-viable 4 (F Viable) 2 (F Non-viable) 3 (M Viable) 0 (M Non- 3 Viable EF viable) 1 Non-viable 0 (F Viable) 1 (F Non-viable) Autumn 2013 0.559 4 (M Viable) 0 (M Non- 4 Viable AL viable) 6 Non-viable 0 (F Viable) 6 (F Non-viable)

Table 4.2 – Gonad viability classifications for P. clava sampled in 2013 from the EF and AL sites

(nEF = 16; nAL = 30). Between-site comparisons were made using Fisher’s Exact test, and associated p-values are presented (α = 0.05). (M = male; F = female; U = sex undetermined)

166

Sampling Season Site n per sex Total n p-value

5 (M Viable) 0 (M Non- 7 Viable EF viable) 3 Non-viable 2 (F Viable) 3 (F Non-viable) Spring 2013 0.685 5 (M Viable) 0 (M Non- 13 Viable AL viable) 3 Non-Viable 8 (F Viable) 3 (F Non-viable) 3 (M Viable) 0 (M Non- 3 Viable EF viable) 7 Non-viable 0 (F Viable) 7 (F Non-viable) Summer 2013 0.266 3 (M Viable) 0 (M Non- 3 Viable AL viable) 2 Non-viable 0 (F Viable) 2 (F Non-viable) 9 (M Viable) 0 (M Non- 9 Viable EF viable) 6 Non-viable 0 (F Viable) 6 (F Non-viable) Autumn 2013 0.350 3 (M Viable) 1 (M Non- 6 Viable AL viable) 1 Non-viable 3 (F Viable) 0 (F Non-viable)

Table 4.3 – Gonad viability classifications for E. dilatata sampled in 2013 from the EF and AL sites (nEF =35; nAL = 28). Between-site comparisons were made using Fisher’s Exact test, and associated p-values are presented (α = 0.05). (M = male; F = female)

167

Female Male Gametogenic Stage Spent/Resorbing 6 Late Atypical Early Atypical

e Developing

v 5 r Late Active

s Early Active

O Inactive

s 4

r 3

n 2

0 Season Spring Summer Autumn Spring Summer Autumn

Figure 4.5 – Gametogenic stages observed in EF P. clava collected and sampled for a biopsy of gonadal fluid in 2013. A total of 11 males and 3 females were identified by biopsy analysis. An additional two samples were classified Inactive however the complete absence of identifiable tissue precluded gender determination.

168

Female Male Gametogenic Stage Spent/Resorbing 9 Early Atypical Mature 8

d Developing

v Late Active

e 7

O 6

n 5

O 4

n 3

C 2

0 Season Spring Summer Autumn Spring Summer Autumn

Figure 4.6 – Gametogenic stages observed in AL P. clava collected and sampled for a biopsy of gonadal fluid in 2013. A total of 9 males and 21 females were sampled for gametogenic analysis.

169

Female Male Gametogenic Stage 9 Spent/Resorbing Late Atypical 8 Early Atypical

e Mature

e 7 Late Active

s 6

n 5

n 3

o C 2

0 Season Spring Summer Autumn Spring Summer Autumn

Figure 4.7 - Gametogenic stages observed in EF E. dilatata collected and sampled for a biopsy of gonadal fluid in 2013. A total of 16 male and 18 female biopsy samples were collected for gametogenic analysis at EF.

170

Female Male Gametogenic Stages 12 Spent/Resorbing Late Atypical Early Atypical 10 Mature Developing Late Active 8

o 6

0 Season Spring Summer Autumn Spring Summer Autumn

Figure 4.8 - Gametogenic stages observed in AL E. dilatata collected and sampled for a biopsy of gonadal fluid in 2013. A total of 12 male and 16 female biopsy samples were collected for gametogenic analysis at AL.

171

12 Expected Observed

s 8

f 6

u 4

0 Female Male

Figure 4.9 – Expected and observed number of P. clava of each sex positively identified at EF

(total n = 21).

172

25 Expected Observed

g 15

b 10

0 F M

Figure 4.10 – Expected and observed number of P. clava of each sex positively identified at AL

(total n = 36). Observed male-to-female ratio (11:25) differed significantly from 1:1.

173

20 Expected Observed

f 10

N 5

0 F M

Figure 4.11 – Expected and observed number of E. dilatata of each sex positively identified at EF

(total n = 29). No significant departure from the expected 1:1 ratio was detected.

174

14 Expected Observed

e 8

V 6

0 F M

Figure 4.12 – Expected and observed number of E. dilatata of each sex positively identified at AL

(total n = 24). No significant departure from the expected 1:1 ratio was detected.

175

EEQ (ng Max E2 Equiv. Conc. Deployment Period Site E2/POCIS) (nM)

EF ND ND* October 2011 – December 2011 AL 0.040 7.166E-2*

EF 0.261 2.986E-1* April 2012 – June 2012 AL 0.304 3.348E-1*

EF 0.438 5.500E-1* June 2012 – August 2012 AL 0.260 2.963E-1*

EF NA 3.231** October 2012 – May 2013 AL ND ND**

EF NA 3.881** May 2013 – July 2013 AL NA 3.357**

EF NA 2.257** July 2013 – September 2013 AL NA 7.202**

Table 4.4 – POCIS estrogenicity values as determined by the YES assay. EEQ values are presented for samples for which an EC20 value could be calculated. Otherwise, the E2 standard concentration equivalent to the maximum observed extract response is provided. Samples that did not produce a response above the upper 99% confidence level of the negative control are presented as ND (no detect), while NA is listed in place of an EEQ for samples that exhibited non-sigmoidal YES assay response curves. *10 µL extract assayed. **200 µL extract assayed.

176

Initially Initially Response Avg (%) Organism Trial Glochidia Open Closed Failure Viability

1 79 76 3 2

AL-1 2 49 44 5 2 86.5

3 25 22 3 2

1 30 29 1 0

AL-2 2 33 33 0 0 97.9

3 33 32 1 0

Table 4.5 – Viability (%) of mature glochidia collected from AL P. clava (n = 2) in early July, 2012.

177

Chapter 5: Conclusion

The major declines that have been observed for unionid populations around the globe, but particularly in North America, where the historical range and past diversity of this group is tremendous in comparison with that seen today, provided the major impetus to conduct the work that comprises this study. Here, a nonlethal biopsy procedure, typically used to determine sex in non-sexually dimorphic unionid species, was utilized in the development, application, and validation of a method to assess gametogenesis and reproductive condition in populations of the unionid species E. complanata located in the Cacapon River (WV). This method was then utilized in the comparative study of two Cacapon River E. complanata populations, located in areas primarily characterized by either agricultural land use or forest. Possible correlations between the reproductive endpoints assessed for these organisms, their proximity to agricultural activity in the region, and the apparent presence or absence of chemical(s) in surface waters having estrogenic activity that has been previously linked to endocrine disruptive effects in a variety of aquatic species were also investigated. Finally, this method was utilized as part of an approach to assess the reproductive success or failure of a population of the federally endangered clubshell, P. clava and proved effective in gaining critical information about the level at which reproduction may be impaired or impeded within the complex reproductive cycle of this unionid species.

Methods developed for application of the nonlethal biopsy procedure in gametogenic assessment in E. complanata consistently provided gametogenic stage categorizations that aligned with those generated using the typical, and lethal, histological approach. Agreement between stages assigned using each of these methods was strong for both male and female

178 individuals of this species during both spring and summer sampling seasons. The method, however, was more successful for male than female specimen, and a number of procedural limitations were evident. Gamete development among unionids, regardless of sex, begins with growth and development of cell stages that retain a physical attachment to acinar walls within the gonad. Data here suggest that the biopsy procedure may be less effective at collecting these attached cell stages than it is for collecting cell stages found unattached within the acinar lumen.

For male unionids, the only cell stage retaining an attachment to the acinar wall is that of the spermatogonium, which is recognized as the initial stage of spermatogenesis. However, the remaining developmental stages of spermatogenesis, including spermatocytes, spermatids, and mature spermatozoa, are all unbound. Therefore, only these unattached cell stages were included in the development of the biopsy-based methodology for male gametogenic assessment. For female unionids, developing ova typically retain a physical attachment to the acinar wall throughout most of development and maturation. While the biopsy may not collect a fully representative sample of the developing tissues of the female gonad, cells collected and used in development of the method for female gametogenic assessment did consistently predict the histologically determined stage when a three-stage classification system of (1) Developing, (2)

Mature, and (3) Spent/Resorbing was utilized. While other, more specific gametogenic classification schemes can, and have been, proposed for female unionids, the ability to drastically limit, or avoid altogether, the need to sacrifice individuals to conduct this gametogenic assessment of a Unioinid population seems an acceptable tradeoff. The procedure defined in this study, provides an approach in which limited histological examination is required in order to determine species-specific threshold ova area values used to define the three gamete stage classifications (Spent/Atretic, Developing, and Mature). However, this might be avoided if an approach to assessing female gamete development is taken similar to that of Barber and Blake

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(1981) and Haggerty et al. (1995), in which ova areas throughout the reproductive cycle are recorded and relative size differences observed over time are used to gauge development and maturity. This alternative approach may be of particular importance for species most critically effected by the recently observed population declines, and for which major conservation endeavors are currently underway, as information on reproduction and gamete development in these species is often lacking or fully absent. Future research efforts should focus on the validation of this approach for additional unionid species.

Following validation of the biopsy procedure for gametogenic assessment of E. complanata in the Cacapon River (WV), this method, among others, was used to compare two populations of this species, one characterized by a proximity to significant agricultural activity, and the other located in an area dominated by forest. Data collected support the finding of a degree of reproductive impairment, particularly for some male individuals, in the agriculture- based population, as well as provide evidence of suppressed growth among individuals at this site.

Additionally, while not statistically significant, qualitative analysis of females sampled suggest some increase in organisms exhibiting a Spent/Resorbing condition at times throughout the reproductive cycle that should see gamete maturation in anticipation of spawn. These observations, particularly for male E. complanata were similar to effects observed for E. dilatata following exposure to varying levels of 17β-estradiol, as these individuals experienced depressed gamete development in response to E2 exposure, relative to control and regardless of dose. In light of these findings, the verified presence of some chemical(s) in surface waters of the Cacapon

River that exhibit estrogenic activity, as determined by YES analysis of POCIS extracts collected from samplers deployed at each site, cannot be discounted (nor can it be verified) as a factor in the developmental effects observed for organisms in this field study.

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Finally, as was proposed earlier, the development and validation of an approach to gametogenic assessment in unionid mussels might prove highly beneficial for species considered threatened or endangered and at particular risk of extinction. While conservation efforts are likely most effectively guided by species-specific information, often the ability to collect such information is greatly limited, as methods, such has been the case historically for assessing gametogenesis and general reproductive development in unionids, often require significant stress to organisms or, at worst, organism sacrifice. These data cannot be collected for highly sensitive species due to these methodological limitations, however, the ability to gauge such parameters without significant stress and in the absence of organism sacrifice is made possible through the biopsy technique, and in this study was applied in the study of one such species as part of a larger investigation into reproductive failure in an apparently declining population of P. clava located in the East Fork West Branch St. Joseph River (MI). Sampling of this population during the course of this study provided evidence of successful recruitment in this small population of P. clava in recent years. However, assessment of gametogenic tissues collected using the biopsy approach suggest a high degree of gametogenic variability, among males in this population. Though sample sizes upon which these conclusions are drawn were small, similar sampling of a “reference” population failed to demonstrate such variability, and provided support of a typical gametogenic cycle in this robust and reproductively active P. clava population, in which rapid development and maturation of gametes precedes spawn during spring to late summer months.

In light of the findings of this work, future conservation efforts and ecotoxicological research on unionid mussels might benefit from a continued focus on development and utilization of nonlethal methods of assessment. A specific recommendation is made for future research in freshwater bivalve ecotoxicology to attempt to utilize nonlethal collection of gonadal tissues during chronic adult exposure studies, in order to investigate effects of exposure to known or

181 suspected EDCs on gametogenesis and gamete viability throughout the reproductive cycle. In addition, whenever possible, further validation of the biopsy technique in other unionid species is recommended by testing agreement between biopsy-based gametogenic staging and histological gametogenic assessment.

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References

Alvarez, D.A. 2010. Guidelines for the use of the semipermeable membrane device (SPMD) and the polar organic chemical integrative sampler (POCIS) in environmental monitoring studies. Ch. 4 of Section D, Water Quality Book 1, Collection of Water Data by Direct Measurement. USGS.

Alvarez, D.A., Cranor, W.L., Perkins, S.D., Randal, C.C., Smith, S.B. 2008. Chemical and Toxicological Assessment of Organic Contaminants in Surface Water Using Passive Samplers. J. Environ. Qual. 37: 1024-1033.

Amyot, J.P., Downing, J.A. 1998. Locomotion in Elliptio complanata (Mollusca: Unionidae): a reproductive function? Freshwater Biology. 39: 351-358.

Anthony, J.L., Kesler, D.H., Downing, W.L., Downing, J.A. 2001. Length-specific growth rates in freshwater mussels (Bivalvia: Unionidae): extreme longevity or generalized growth cessation? Freshwater Biology, 46: 1349-1359.

Augspurger, T., Dwyer, F.J., Ingersoll, C.G., Kane, C.M. 2007. Advances and opportunities in assessing contaminant sensitivity of freshwater mussel (Unionidae) early life stages. Environmental Toxicology and Chemistry 26: 2025.

Barber, B.J. 1996. Gametogenesis of eastern oysters, Crassostrea virginica (Gmelin, 1791), and pacific oysters, Crassostrea gigas (Thunberg, 1793) in disease-endemic lower Chesapeake Bay. Journal of Shellfish Research. 15(2): 285-290.

Barber, B.J., Blake, N.J. 1981. Energy storage and utilization in relation to gametogenesis in Argopecten irradians concentricus (Say). J. exp. Mar. Biol. Ecol. 52: 121-134.

Bauer, G. 1992. Variation in the life span and size of the freshwater pearl mussel. Journal of Animal Ecology, 61: 425-436.

Beasley, C.R., Tury, E., Vale, W.G., Tagliaro, C.H. 2000. Reproductive cycle, management and conservation of Paxyodon syrmatophorus (Bivalvia: Hyriidae) from the Tocantins River, Brazil. J. Moll. Stud. 66: 393-402.

Berg, D.J., Haag, W.R., Guttman, S.I., Sickel, J.B. 1995. Mantle biopsy: a technique for nondestructive tissue-sampling of freshwater mussels. J.N. Am Benthol Soc. 14:577-581.

Blazer, V.S., Iwanowicz, L.R., Iwanowicz, D.D., Smith, D.R., Young, J.A., Hedrick, J.D., Foster, S.W., Reeser, S.J. 2007. Intersex (Testicular Oocytes) in Smallmouth Bass from the Potomac River and Selected Nearby Drainages. Journal of Aquatic Animal Health 19 : 242–253.

183

Bogan, A.E., Roe, K.J. 2008. Freshwater bivalve (Unioniformes) diversity, systematics, and evolution: status and future directions. J.N. Am. Benthol. Soc. 27(2):349-369.

Breton, S., Doucet-Beaupre, H., Stewart, D.T., Hoeh, W.R., Blier, P.U. 2007. The unusual system of doubly uniparental inheritance of mtDNA: isn’t one enough? Trends in Genetics, 23: 465- 474.

Bringolf, R.B., Cope, W.G., Eads, C.B. 2007a. Acute and Chronic toxicity of technical-grade pesticides to glochidial and juveniles of freshwater mussels (Unionidae). Environmental Toxicology and Chemistry, 26(10): 2086-2093.

Bringolf, R.B., Cope, W.G., Mosher, S. 2007b. Acute and Chronic toxicity of glyphosate to glochidial and juveniles of Lampsilis siliquoidea (Unionidae). Environmental Toxicology and Chemistry, 26(10): 2094-2100.

Bringolf, R.B., Cope, W.G., Barnhart, M.C. 2007c. Acute and Chronic toxicity of pesticide formulations (atrazine, chlorpyrifos, and permethrin) to glochidial and juveniles of Lampsilis siliquoidea. Environmental Toxicology and Chemistry, 26(10): 2101-2107.

Buckley, J.A. 2010. Quantifying the antiestrogen activity of wastewater treatment plant effluent using the yeast estrogen screen. Environmental Toxicology and Chemistry, 29(1): 73-78.

Cek, S., Sereflisan, H. 2011. The Gametogenic Cycle of Leguminaia whaetleyi (Lea, 1862) in Lake Golbasi, Turkey (Bivalvia: Unionidae). Journal of Experimental Zoology. 315: 30-40.

Chambers, D.B., Leiker, T.J. 2006. A Reconnaissance for Emerging Contaminants in the South Branch Potomac River, Cacapon River, and Williams River Basins, West Virginia, April- October 2004. U.S. Geological Survey Open-File Report 2006-1393, 23 p. http://pubs.usgs.gov/of/2006/1393

Chesman, B.S., Langston, W.J. 2006. Intersex in the clam Scrobicularia plana: a sign of endocrine disruption in estuaries? Biology Letters, 2(3): 420-422.

Christian, A.D., Harris, J.L. 2008. An introduction to directions in freshwater mollusk conservation: molecules to ecosystems. J.N. Am. Benthol. Soc. 27(2):345-348.

Ciocan, C.M., Cubero-Leon, E., Minier, C., Rotchell, J.M. 2011. Identification of Reproduction- Specific Genes Associated with Maturation and Estrogen Exposure in a Marine Bivalve Mytilus edulis. PLoS One. 6(7): e22326. doi: 10.1371/journal.pone.0022326.

Colucci, M.S., Bork, H., Topp, E. 2001. Persistence of estrogenic hormones in agricultural soils: I. 17Beta-estradiol and estrone. J. Environ. Qual. 30: 2070-2076.

Combalbert, S., Hernandez-Raquet, G. 2010. Occurrence, fate, and biodegradation of estrogens in sewage and manure. Appl Microbiol Biotechnol. 86: 1671-1692.

Constantz, G., Ailes, N., Malakoff, D. 2005. Portrait Of A River: The Ecological Baseline of the Cacapon River. 32p. http://www.cacaponinstitute.org/PDF/Publications/Portrait%20of%20a%20River.pdf

184

Croll, R.P., Wang, C. 2007. Possible roles of sex steroids in the control of reproduction in bivalve molluscs. Aquaculture. 272: 76-86.

Dillon, R.T. Jr. 2004. The Ecology of Freshwater Molluscs. Cambridge University Press, Cambridge, UK.

Dinamani, P. 1974. Reproductive cycle and gonadial changes in the New Zealand rock oyster Crassostrea glomerate. New Zealand Journal of Marine and Freshwater Research. 8(1):39- 65.

Downing, J.A., Rochon, Y., Perusse, M., Harvey, H. 1993. Spatial aggregation, body size, and reproductive success in the freshwater mussel Elliptio complanata. J.N. Am. Benthol. Soc. 12(2): 148-156.

Du, B., Haddad, S.P., Luek, A., Scott, W.C., Saari, G.N., Kristofco, L.A., Connors, K.A., Rash, C., Rasmussen, J.B., Chambliss, C.K., Brooks, B.W. 2014. Bioaccumulation and trophic dilution of human pharmaceuticals across trophic positions of an effluent-dependent wadeable stream. Phil. Trans. R. Soc. B. 369: 20140058. http://dx.doi.org/10.1098/rstb.2014.0058.

Eldridge, J.C., Stevens, J.T., Breckenridge, C.B. 2008. Atrazine interaction with estrogen expression systems. Rev. Environ. Contam. Toxicol. 196: 147-160.

Elrashidi, M.A., Seybold, C.A., Wysocki, D.A., Peaslee, S.D., Ferguson, R., West, L.T. 2008. Phosphorus in runoff from two watersheds in Lost River Basin, West Virginia. Soil Science. 173(11): 792-806.

Farris, J.L. and Van Hassel, J.H. 2007. Freshwater Bivalve Ecotoxicology. CRC Press – Taylor and Francis Group, New York, in collaboration with the Society of Environmental Toxicology and Chemistry (SETAC), Florida.

Finlay-Moore O., Hartel, P.G., Cabrera, M.L. 2000. 17β-Estradiol and testosterone in soil and runoff from grasslands amended with broiler litter. J Environ. Qual. 29:1604-1611.

Frye, C., Bo, E., Calamandrei, G., Calzà, L., Dessi-Fulgheri, F., Fernández, M., Fusani, L., Kah, O., Kajta, M., Le Page, Y., Patisaul, H.B., Venerosi, A., Wojtowicz, A.K., Panzica, G.C. 2013. Endocrine Disrupters: A Review of Some Sources, Effects, and Mechanisms of Actions on Behavior and Neuroendocrine Systems. J. Neuroendocrinol. 24(1): 144-159.

Gagné, F., Blaise, C. 2003. Effects of municipal effluents on serotonin and dopamine levels in the freshwater mussel Elliptio complanata. Comparative Biochemistry and Physiology Part C 136 : 117-125.

Gagné, F., Marcogliese, D.J., Blaise, C., Gendron, A.D. 2001. Occurrence of Compounds Estrogenic to Freshwater Mussels in Surface Waters in an Urban Area. Environ Toxicol. 16 (3) : 260- 268.

Garner, J.T., Haggerty, T.M., Modlin, R.F. 1999. Reproductive cycle of Quadrula metanevra (Bivalvia: Unionidae) in the Pickwick Dam tailwater of the Tennessee River. American Midland Naturalist. 141: 277-283.

185

Giusti, A., Joaquim-Justo, C. 2013. Esterification of vertebrate like steroids in molluscs: A target of endocrine disruptors? Comparative Biochemistry and Physiology, Part C, 158: 187-198.

Gordon, M. E., Smith, D. G. 1990. Autumnal reproduction in Cumberlandia monodonta (Unionoidea: Margaritiferidae). Transactions of the American Microscopical Society. 109: 407–411.

Gourmelon, A., Ahtiainen, J. 2007. Developing test guidelines on invertebrate development and reproduction for the assessment of chemicals, including potential endocrine active substances – The OECD Perspective. Ecotoxicology. 16: 161 – 167.

Grande, C., Araujo, R., Ramos, M.A. 2001. The gonads of Margaritifera auricularia (Spengler, 1793) and M. margaritifera (Linnaeus, 1758) (Bivalvia: Unionoidea). Journal of Molluscan Studies. 67: 27–35.

Gustafson, L.L., Stoskoph, M.K., Bogan, A.E., Showers, W., Kwak, T.J., Hanlon, S., Levine, J.F. 2005. Evaluation of a nonlethal technique for hemolymph collection in Elliptio complanata, a freshwater bivalve (Mollusca: Unionidae). Diseases of Aquatic Organisms. 65: 159-165.

Haag, W.R., Stanton, J.L. 2003. Variation in fecundity and other reproductive traits in freshwater mussels. Freshwater Biology. 48: 2118-2130.

Haag, W.R., Williams, J.D. 2014. Biodiversity on the brink: an assessment of conservation strategies for North American freshwater mussels. Hydrobiologia. 735: 45-60.

Haggerty, T.M., Garner, J.T., Patterson, G.H., Jones Jr., L.C. 1995. A quantitative assessment of the reproductive biology of Cyclonaias tuberculate (Bivalvia: Unionidae) Can. J. Zool. 73: 83- 88.

Hanselman, T.A., Graetz, D.A., Wilkie, A.C. 2003. Manure-Borne Estrogens as Potential Environmental Contaminants: A Review. Environmental Science & Technology 37 (24): 5471-5478.

Hardege, J.D., Duncan, J., Ram, J.L. 1997. Tricyclic Antidepressants Suppress Spawning and Fertilization in the Zebra Mussel Dreissena polymorpha. Comp. Biochem. Physiol. 118C (1): 59-64.

Heard W.H. 1979. Hermaphroditism in Elliptio (Pelecypoda: Unionidae). Malacological Review, 12: 21-28.

Henley, W.F. 2002. Evaluation of diet, gametogenesis, and hermaphroditism in freshwater mussels (Bivalvia: Unionidae). Ph.D. Dissertation. Virginia Polytechnic Institute and State University.

Henley, W.F., Neves, R.J., Caceci, R., Saacke, R.G. 2007. Anatomical descriptions and comparison of the reproductive tracts of Utterbackia imbecillis and Villosa iris (Bivalvia: Unionidae). Invertebrate Reproduction and Development. 50: 1–12.

186

Ingersoll, C.G., Kernaghan, N.J., Gross, T.S., Bishop, C.D., Wang, N., Roberts, A. 2007. Laboratory toxicity testing with freshwater mussels. In Freshwater Bivalve Ecotoxicology, CRC Press, New York, NY; p. 95-134.

Janer, G., Lavado, R., Thibaut, R., Porte, C. 2005. Effects of 17β-estradiol exposure in the mussel Mytilus galloprovincialis: A possible regulating role for steroid acyltransferases. Aquatic Toxicology, 75: 32-42.

Janer, G., Porte, C. 2007. Sex steroids and potential mechanisms of non-genomic endocrine disruption in invertebrates. Ecotoxicology. 16: 145-160.

Jarosova, B., Blaha, L., Vrana, B., Randak, T., Grabic, R., Biesy, J.P., Hilscherova, K. 2012. Changes in concentrations of hydrophilic organic contaminants and endocrine-disrupting potential downstream of small communities located adjacent to headwaters. Environment International, 45: 22-31.

Jirka, K. J., Neves, R. J. 1992. Reproductive biology of four species of freshwater mussels (Mollusca: Unionidae) in the New River, Virginia and West Virginia. Journal of Freshwater Ecology. 7: 35–44.

Jobling, S. Casey, D. Rodgers-Gray, T. Oehlmann, J., Schulte-Oehlmann, U., Pawlowski, S. Baunbeck, T. Turner, A.P., Tyler, C.R. 2004. Comparative responses of molluscs and fish to environmental estrogens and an estrogenic effluent. Aquatic Toxicology 66: 207-222.

Jones, H.A., Simpson, R.D., Humphrey, C.L. 1986. The reproductive cycles and glochidial of freshwater mussels (Bivalvia: Hyriidae) of the Macleay River, Northern New South Wales, Australia. Malacologia. 27: 185-202.

Karlsson, S., Larsen, B.M., Eriksen, L., Hagen, M. (2013). Four methods of nondestructive DNA sampling from freshwater pearl mussels Margaritifera margaritifera L. (Bivalvia: Unionoida). Freshwater Science. 32(2): 525 – 530.

Kat, P.W. 1983. Sexual selection and simultaneous hermaphroditism among the Unionidae (Bivalvia: Mollusca). J. Zool., Lond. 201: 395-416.

Keller, A., Lydy, M., Ruessler, D.S. 2007. Unionid mussel sensitivity to environmental contaminants. In Freshwater Bivalve Ecotoxicology, CRC Press, New York, NY; p. 151-167.

Kesler, D.H., Newton, T.J., Green, L. 2007. Long-term monitoring of growth in the Eastern Elliptio, Elliptio complanata (Bivalvia: Unionidae), in Rhode Island: a transplant experiment. J.N. Am. Benthol. Soc. 26(1): 123-133.

Khanal, S.K., Xie, B., Thompson, M.L. 2006. Fate, transport, and biodegradation of natural estrogens in the environment and engineered systems. Environmental Science and Technology, 40(21): 6537-6546.

Klotz, D.M., Beckman, B.S., Hill, S.M., McLachlan, J.A., Walters, M.R., Arnold, S.F. 1996. Identification of environmental chemicals with estrogenic activity using a combination of in vitro assays. Environ. Health Perspect. 104(10): 1084-1089.

187

Labadie, P., Peck, M., Minier, C., Hill, E.M. 2007. Identification of the steroids fatty acid ester conjugates formed in vivo in Mytilus edulis as a result of exposure to estrogens. Steroids, 72: 41-49.

Landis, J.R., Koch, G.G. 1977. The measurement of observer agreement for categorical data. Biometrics. 33(1): 159-174.

Landis, W.G., Yu, M. 2004. Introduction to Environmental Toxicology. Ed. 3. Lewis Publishers – A CRC Press Company, New York.

Leonard, J.A., Cope, W.G., Barnhart, M.C., Bringold, R.B. 2014a. Metabolomic, behavioral, and reproductive effects of the synthetic estrogen 17α-ethinylestradiol on the Unionid mussel Lampsilis fasciola. Aquatic toxicology. 150: 103-116.

Leonard, J.A., Cope, W.G., Barnhart, M.C., Bringolf, R.B. 2014b. Metabolomic, behavioral, and reproductive effects of the aromatase inhibitor fadrozole hydrochloride on the Unionid mussel Lampsilis fasciola. General and Comparative Endocrinology. 206: 213-226.

Liu, W., Li, Q., Kong, L. 2008. Estradiol-17 and testosterone levels in the cockle Fulvia mutica during the annual reproductive cycle. New Zealand Journal of Marine and Freshwater Research. 42: 411-424.

Matteson, M.R. 1948. Life History of Elliptio complanatus (Dillwyn, 1817). American Midland Naturalist, 40(3): 690-723.

McElwain, A., Bullard, S.A. 2014. Histological atlas of freshwater mussels (Bivalvia: Unionidae): Villosa nedulosa (Ambleminae: Lampsilini), Fusconaia cerina (Ambleminae: Pleurobemini), and Strophitus connasaugaensis (Unioninae: Anodontini). Malacologia, 57(1): 99 – 239.

Naimo, T.J., Damschen, E.D., Rada, R.G., Monroe, E.M. 1998. Nonlethal evaluation of the physiological health of Unionid mussels: methods for biopsy and glycogen analysis. J.N. Am Benthol Soc. 17(1): 121-128.

Nelson J., Bishay F., van Roodselaar A. Ikonomou M., Law F.C.P. 2007. The use of in vitro bioassays to quantify endocrine disrupting chemicals in municipal wastewater treatment plants effluents. Sci Total Environ. 374:80–90.

Newton, T.J., Bartsch, M.R. 2007. Lethal and sublethal effects of ammonia to juvenile Lampsilis mussels (Unionidae) in sediment and water-only exposures. Environmental Toxicology and Chemistry, 26(10): 2057-2065.

Nentwig, G. 2007. Effects of Pharmaceuticals on aquatic invertebrates. Part II: The antidepressant drug Fluoxetine. Arch. Environ. Contam. Toxicol. 52: 163-170.

Nichols, D.J., Daniel, T.C., Edwards, D.R., Moore, P.A. Jr., Pote, D.H. 1998. Use of grass filter strips to reduce 17 beta-estradiol in runoff from fescue-applied poultry litter. J. Soil Water Conserv. 53, 74-77.

188

Nichols, D.J., Daniel, T.C., Moore, P.A. Jr., Edwards, D.R., Pote, D.H. 1997. Runoff of Estrogen hormone 17β-estradiol from poultry litter applied to pasture. Journal of Environmental Quality 26 (4): 1002-1006.

O’Dee, S.H., Watters, G.T. 1998. New or confirmed host identifications for ten freshwater mussels. Proceedings of the Conservation, Captive Care, and Propagation of Freshwater Mussels Symposium. 77-82.

Ortiz-Zarragoitia, M., Cajaraville, M.P. 2006. Biomarkers of exposure and reproduction-related effects in mussels exposed to endocrine disruptors. Arch. Environ. Contam. Toxicol. 50: 361–369.

Ortiz-Zarragoitia, M., Cajaraville, M.P. 2010. Intersex and oocyte atresia in a mussel population from the Biosphere’s Reserve of Urdaibai (Bay of Biscay). Ecotoxicology and Environmental Safety, 73: 693-701.

Peck, M.R., Labadie, P., Minier, C., Hill, E.M. 2007. Profiles of environmental and endogenous estrogens in the zebra mussel Dreissena polymorpha. Chemosphere. 69: 1-8.

Peterson, E.W., Davis R.K., Orndorff, H.A. 2000. 17β-estradiol as an indicator of animal waste contamination in mantled karst aquifers. J. Environ. Qual. 29: 826-834.

Philips, S.W. 2005. U.S. Geological Survey Chesapeake Bay Science Plan, 2006 – 2011. Open-File Report 2005-1440.

Rastall, A.C. 2004. The development of a biomimetic approach to the detection and identification of anthropogenic estrogen receptor agonists in surface waters. Ph.D. Dissertation. The Open Univ., United Kingdom.

Rastall, A.C., Neziri, A., Vukonvic, Z., Jung, C., Mijovic, S., Hollert, H., Nikcevic, S., Erdinger, L. 2004. The Identification of Readily Bioavailable Pollutants in Lake Shkodra/Skadar Using Semipermeable Membrane Devices (SPMDs), Bioassays and Chemical Analysis. Environ Sci & Pollut Res. 11: 240-253.

Regoli, L., Chan, H.M., de Lafontaine, Y., Mikaelian, I. 2001. Organotins in zebra mussels (Dreissena polymorpha) and sediments of the Quebec City Harbour area of the St. Lawrence River. Aquatic Toxicology 53 (2) : 115-126.

Ricciardi, A., Rasmussen, J.B. 1999. Extinction rates of North American freshwater fauna. Conservation Biology. 13(5): 1220-1222

Routledge, E.J., Sumpter, J.P. 1996. Estrogenic activity of surfactants and some of their degradation products assessed using a recombinant yeast screen. Environmental Toxicology and Chemistry. 15(3): 241-248.

Roznere, I., Watters, G.T., Wolf, B.A., Daly, M. 2014. Nontargeted metabolomics reveals biochemical pathways altered in response to captivity and food limitation in the freshwater mussel Amblema plicata. Comparative Biochemistry and Physiology, Part D, 12: 53-60.

189

Saha, S., Layzer, J.B. 2008. Evaluation of a nonlethal technique for determining sex of freshwater mussels. J.N. Am. Benthol. Soc. 27(1): 84-89.

Scott, A.P. 2012. Do mollusks use vertebrate sex steroids as reproductive hormones? Part I: Critical appraisal of the evidence for the presence, biosynthesis and uptake of steroids. Steroids. 77: 1450-1468.

Scott, A.P. 2013. Do mollusks use vertebrate sex steroids as reproductive hormones? II. Critical review of the evidence that steroids have biological effects. Steroids. 78: 268-281.

Sereflisan, H., Cek, S., Sereflisan, M. 2013. The reproductive cycle of Potomida littoralis (Cuvier, 1798) (Bivalvia: Unionidae) in Lake Golbasi, Turkey. Pakistan Journal of Zoology. 45(5).

Shepardson, S.P., Heard, W.H., Breton, S., Hoeh, W.R. 2012. Light and transmission electron microscopy of two spermatogenic pathways and unimorphic spermatozoa in Venustaconcha ellipsiformis (Conrad, 1836) (Bivalvia: Unionoida). Malacologia. 55(2): 263-284.

SJRWI. 2012. St. Joseph River Watershed Initiative: Water Quality Monitoring Report 2011. http://www.sjrwi.org/sites/default/files/docs/publications/2011%20Report.pdf

Smith, D.G. 1978. Biannual gametogenesis in Margaratifera margaritifera (L.) in northeastern North America. Bulletin of the American Malacological Union. 49-53.

Smith, D.G. 1988. Notes on the biology and morphology of Margaritifera hembeli (Conrad, 1838) (Unionacea: Margaritiferidae). The Nautilus. 102: 159-163.

Smith, D.G., Lang, B.K., Gordon, M.E. 2003. Gametogenic cycle, reproductive anatomy, and larval morphology of Popenaias popeii (Unionoida) from the Black River, New Mexico. Southeastern Naturalist. 48: 333–340.

Sokal, R.R., Rohlf, F.J. 1998. Biometry – The Principles and Practice of Statistics in Biological Research. Third Ed. W.H. Freeman and Company, New York, NY.

Strayer, D.L., Downing, J.A., Haag, W.R., King, T.L., Layzer, J.B., Newton, T.J., Nichols, S.J. 2004. Changing perspectives on Pearly mussels, North America’s most imperiled animals. BioScience 54(5): 429-439.

Strayer, D.L., Malcom, M. 2012. Causes of recruitment failure in freshwater mussel populations in southeastern New York. Ecological Applications. 22(6): 1780 - 1790.

Strayer, D.L., Smith, D.R. 2003. A guide to sampling freshwater mussel populations. American Fisheries Society, Monograph 8: 110pp.

Sumpter, J.P. 2005. Endocrine Disrupters in the Aquatic Environment: An Overview. Acta hydrochim. hydrobiol. 33(1): 9-16.

TNNMCC. 1998. National Strategy for the Conservation of Native Freshwater Mussels. Prepared by The National Native Mussel Conservation Committee. Journal of Shellfish Research, 17(5): 1419-1428.

190

USFWS. 1994. U.S. Fish and Wildlife Service. Clubshell (Pleurobema clava) and northern riffleshell (Epioblasma torulosa rangiana) recovery plan. Hadley, Massachusetts. 68 pp.

USFWS. 2015. ECOS – Environmental Conservation Online System: Listed Animals. http://ecos.fws.gov/tess_public/reports/ad-hoc-species- report?kingdom=V&kingdom=I&status=E&status=T&status=EmE&status=EmT&status=E XPE&status=EXPN&status=SAE&status=SAT&mapstatus=3&fcrithab=on&fstatus=on&fsp ecrule=on&finvpop=on&fgroup=on&header=Listed+Animals

Van Der Schalie, H. 1970. Hermaphroditism Among North American Freshwater Mussels. Malacologia, 10(1): 93-112.

Van Der Schalie, H., Van Der Schalie, A. 1963. Occasional papers of the museum of zoology University of Michigan.

Villella, R.F., Smith, D.R. 2005. Two-phase sampling to estimate river-wide populations of freshwater mussels. J.N. Am. Benthol. Soc. 24(2): 357-368.

Wang, C., Croll, R.P. 2004. Effects of sex steroids on gonadal development and gender determination in the sea scallop, Placopecten magellanicus. Aquaculture, 238: 483-498.

Wang, N., Augspurger, T., Barnhart, M.C., Bidwell, J.R., Cope, W.G., Dwyer, F.J., Geis, S., Greer, I.E., Ingersoll, C.G., Kane, C.M., May, T.W., Neves, R.J., Newton, T.J., Roberts, A.D., Whites, D.W. 2007. Intra-and interlaboratory variability in acute toxicity tests with glochidial and juveniles of freshwater mussels (Unionidae). Environmental Toxicology and Chemistry, 26(10): 2029-2035.

Wang, N., Erickson, R.J., Ingersoll, C.G. 2008. Influence of pH on the acute toxicity of ammonia to juvenile freshwater mussels (fatmucket, Lampsilis siliquoidea). Environmental Toxicology and Chemistry, 27(5): 1141-1146.

Watermann, B., Thomsen, A., Kolodzey, H., Daehne, B., Meemken, M., Pijanowska, U., Liebezeit, G. 2008. Histopathological lesions of molluscs in the harbor of Norderney, Lower Saxony, North Sea (Germany). Helgol Mar Res. 62: 167-175.

Watters, G.T. 1988. A survey of the freshwater mussels of the St. Joseph River system, with emphasis on the federally endangered White Cats Paw Pearly Mussel. Final Report to the Division of Fish and Wildlife, Indiana Department of Natural Resources. 127pp.

Watters, G.T. 2007. A Brief Look at Freshwater Mussel (Unionacea) Biology. In Freshwater Bivalve Ecotoxicology, CRC Press, New York, NY; p. 51-64.

Weaver, L.R., Pardue, G.B., Neves, R.J. 1991. Reproductive biology and fish hosts of the Tennessee Clubshell, Pleurobema oviforme (Mollusca: Unionidae) in Virginia. American Midland Naturalist. 126(1): 82-89.

Williams, J.D., Warren Jr, M.L., Cummings, K.S., Harris, J.L, Neves, R.J. 1993. Conservation Status of Freshwater Mussels of the United States and Canada. Fisheries, 18(9): 6 – 23.

191

Winslow, M.L. 1918. Pleurobema clava (Lam.) and Planorbis dilatatus buchanensis (Lea) in Michigan. Occasional Papers of the Museum of Zoology, University of Michigan, 51:1-4.

Yang, R. Li, N., Rao, K., Ma, M., Wang, Z. 2015. Combined action of estrogen receptor agonists and antagonists in two-hybrid recombinant yeast in vitro. Ecotoxicology and Environmental Safety, 111: 228-235.

Yeager, B.L., Neves, R.J. 1986. Reproductive cycle and fish hosts of the Rabbit’s Foot mussel, Quadrula cylindrical strigillata (Molusca: Unionidae) in the Upper Tennessee river drainage. American Midland Naturalist. 116(2): 329-340.

Yokely, Jr. P. 1972. The life history of Pleurobema cordatum (Rafinesque 1820) (Bivalvia: Unionacea). Malacologia. 11: 351-364.

Zale, A.V., Neves, R.J. 1982. Reproductive biology of four freshwater mussel species (Mollusca: Unionidae) in Virginia. Freshwater Invertebrate Biology. 1:17-28.

192