Thesis Title Here

Home , Amblema, Amblema plicata, Freshwater pearl mussel

Methods in health assessment of freshwater mussels, Amblema plicata and Quadrula spp.

Thesis

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

K. Hope Valentine, D.V.M.

Graduate Program in Veterinary Preventive Medicine

The Ohio State University

2011

Thesis Committee:

Paivi Rajala-Schultz, Advisor

Barbara Wolfe

Mary Jo Burkhard

K. Hope Valentine

2011

Abstract

Health monitoring that is non-invasive, repeatable, and sensitive is a critical need for North America’s most endangered animal, the freshwater mussel. Ohio is home to a large proportion of these threatened animals and in response has established a captive flow through facility for research and propagation. Currently there is gap in knowledge of mussel physiology and health requirements in the wild and in captivity, representing an urgent need for health assessment methods. Hemolymph, the circulatory fluid of bivalves, and the cellular portion called hemocytes, are just beginning to be investigated in freshwater mussels. Recent studies have shown that hemolymph can be safely and repeatedly drawn. In addition, a schematic for freshwater mussels’ hemocytes was developed using L-cysteine as a novel anticoagulant. Following trends in hemolymph biochemistry and cellular differentials may be a way to monitor the health of freshwater mussel populations in the wild and in captivity; however, current handling protocols for transport of hemolymph are unknown. This thesis aimed to develop hemolymph handling protocols to preserve cellular integrity and function over time and during transport; to develop baseline biochemical and hematologic reference ranges in a population of wild mussels; and to describe population trends in biochemical and hematological parameters over one year following translocation of animals into captivity.

Due to the marked physiological differences between invertebrate and vertebrate species, handling protocols for hemolymph and cell enumeration cannot be inferred from mammalian based research. Given that the majority of laboratories are 1-4 hours away from the wild locations or captive facility, handling protocols during transport to minimize in-vitro effects were developed. The second chapter of this thesis investigated processing temperatures for hemolymph, the ideal pH for reconstitution of the anticoagulant L-cysteine, and the effects of prolonged exposure to L-cysteine in-vitro.

Towards an optimal method for hemocyte enumeration, cytochalasin B, genistein, and a lower dose of formalin than previously published were evaluated. Total hemocyte count, percentage of viable cells, and presence or absence of cellular debris as an indicator of cell lysis and degranulation were compared for each experiment. Chapter 2 found that exposure of hemocytes to temperatures below 10 degrees Celsius increased cellular aggregation in-vitro and decreased cellular viability. Only formalin treatments were found to increase total cell counts. Treatment of hemocytes with formalin, L-cysteine, cytochalasin B, and genistein resulted in a marked production of cellular debris at one hour post exposure. Untreated hemocytes at ambient temperature can maintain a high level of viability (80-93%) for up to 24 hours. Therefore, hemocytes should be transported without fixative or anticoagulant at ambient temperature. It is recommended that hemocytes only be exposed to L-cysteine (25mg/ml, pH 8.0) just prior to slide preparations for morphological analysis.

Hemolymph chemistry may be a useful non-lethal indicator of bivalve physiological processes once reference ranges are established to differentiate normal

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from abnormal fluctuations in health. Chapter 3 aimed to establish reference ranges for the following measurable components in hemolymph: sodium, chloride, magnesium, phosphorus, potassium, calcium, glucose, and isoenzymes alanine aminotransferase

(ALT), aspartate aminotransferase (AST), and alkaline phosphatase (ALP) in a population of mussels in the wild and follow changes in the biochemistry of hemolymph from the same population translocated into captivity over one year. Hemolymph from forty animals of three species, Amblema plicata, Quadrula quadrula, and Quadrula pustulosa was collected in July of 2008 from the Muskingum River in Devola, Ohio to develop baseline reference ranges. Thirty of those forty animals were translocated into captivity, with nine captive controls. Animals were sampled biweekly for the first month and then quarterly over one year. Significant differences in sodium, potassium, chloride, magnesium, ALT, AST, and ALP were found between genera, A. plicata and Quadrula spp. at baseline (p<0.05). Both genera showed declines in sodium and chloride in the first month in captivity and had marked increases in all electrolyte values in November, five months after transport to captivity. Calcium and glucose values remained steady in the population until the last collection point in June when values in both parameters declined. Phosphorus levels increased in both genera with significantly higher levels seen in A. plicata in February (p<0.05). Losses occurred during the study in greater proportion after winter sampling time points, potentially indicating that sampling during winter quiescence is not recommended. This study indicates that genus differences are present in biochemical values in freshwater mussel hemolymph in the wild and in captivity over time.

In mammalian medicine, a complete blood cell count is a crucial piece of the minimal health database for diagnosis of potential disease or routine health checkups.

Recent development of a standardized method for hemocyte characterization in freshwater mussels has paved the way for application of this technique to monitor freshwater mussels in the wild and in captivity. However, before interpretations of normal and abnormal fluctuations in cell type can take place, reference ranges need to be developed. This study aimed to provide preliminary baseline reference ranges from two common freshwater mussel genera, Amblema and Quadrula in the wild during the peak summer field season, then translocated animals into captivity for monitoring of hemocytology trends in a captivity over one year. Cell differentials were found to be genus-specific at baseline and for the first month in captivity (p< 0.05). Total hemocyte counts between genera differed significantly at 2wks and in November (p <0.10). Four cell types were identified in each genus studied at each time point. Eosinophilic granulocytes predominated in both genera. The proportion of eosinophilic granulocytes in A. plicata ranged from 53 to73%, large agranulocytes from 19 to 41%, basophilic granulocytes from 1 to 6% and small agranulocytes from 1to 3%, compared to Quadrula spp. that had 44-61% eosinophilic granulocytes, 8-27% basophilic granulocytes, 28-40% large agranulocytes, and less than 1 % small agranulocytes. This study provides a foundation for baseline reference ranges for A. plicata and Quadrula spp. and a preliminary understanding of shifts in blood cells in a population of mussels in captivity over one year.

Dedication

This thesis is dedicated to my mentor Dr. Barbara Wolfe who has inspired, encouraged, and pushed me to be and do better whether near or far, and to my parents, Karla Hehl and

Val Valentine, without whom none of this would have been possible.

Acknowledgements

I would like to thank Dr. Paivi Rajala-Schultz, for her pragmatic advice and guidance at every step of development of this degree from the inception of the idea to pursue it to its completion and Dr. Mary Jo Burkhard for her expertise in clinical pathology and willingness to study bivalves even though she is deathly allergic to shellfish.

To Dr. Amanda Nahlik, my dear friend that really pushed me through the final stages of my writing, providing invaluable support and feedback and reading the umpteenth copy of each chapter. To Casey Pollack for letting me use her primo cubicle space and Dr. Mary Kantula, Steve Cline, Teresa McGee, Dixon Flanders, Phil

Kaufmann and Glenn Griffith for their inspiration in the last few weeks of writing. I would also like to thank Sarah Josephine Fannin for making everything more worthwhile and her continuous support to keep going and stay focused.

Finally, I would like to thank Trisha Gibson, Brooke Kelly, Kody Kuehnl, and

Tom Watters from the Columbus Zoo and Aquarium Freshwater mussel Conservation

Research Center for their mussel expertise and dedication to the conservation of these imperiled amazing little creatures. Many thanks to the Morris Animal foundation, the

Ohio Division of Wildlife, the Columbus Foundation and Chemical Abstracts for support of this research.

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Vita

December 1 1979 ...... Born- Raleigh, North Carolina

2002...... B.S. Animal Science and Wildlife

Conservation, University of Massachusetts-

Amherst. Amherst

2007 ...... D.VM., North Carolina State University,

College of Veterinary Medicine, Raleigh

2007-2009 ...... Conservation Medicine Intern, the Wilds,

Cumberland, OH

2008-2010 ...... M.S. Veterinary Preventive Medicine, The

Ohio State University, Columbus

Publications

Valentine, K.H., C.A. Harms , M.B. Cadenas, A.J Birkenheuer, H.S. Marr, R.G Maggi, and E.B. Breitschwerdt. 2007. Bartonella DNA in loggerhead sea turtles (Caretta caretta). Emerging Infectious Diseases. 13(6): 949-950.

Burkhard, M.J., S. Leavell, R.B. Weiss, K. Kuehnl, H. Valentine, G.T.Watters and B.A. Wolfe. Analysis and cytologic characterization of hemocytes from freshwater mussels (Quadrula spp.). Veterinary Clinical Pathology. 38(2): 1-11.

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Oral Presentations

Valentine, K.H, MJ Burkhard, S Leavell, and BA Wolfe. In vitro processing effects on the viability of hemocytes of freshwater mussels (Quadrula spp). International Association of Aquatic Animal Medicine, San Antonio, TX, May 2009.

Valentine, KH, M Abley, R.B. Weiss, and B.A.Wolfe. Salmonella and Campylobacter in exotic hoofstock and carnivores. Ann. Conf. Am. Assoc. Zoo Veterinarians. Los Angeles, CA, October 2008.

Valentine, K.H., B.A. Wolfe, M.J. Burkhard, S. Leavell, R.B. Weiss, K. Kuehnl, and GT Watters. Improving assessment of health and stress in translocated freshwater mussels. Annual Conference of the American Malacological Association, Carbondale, IL, June 2008.

Awards

Post Doctorate Oral Presentation Award, International Association of Aquatic Animal Medicine, 2009, 2nd place.

Fields of Study

Major Field: Veterinary Preventive Medicine

Table of Contents

Abstract ...... ii Dedication ...... vi Acknowledgements ...... vii Vita ...... viii List of Tables ...... xiii List of Figures ...... xvi Chapter 1: Introduction ...... 1 1-1 Introduction 1 Introduction to Unionids 3 References 7 Chapter 2: Processing methods for hemolymph and efforts to reduce hemocyte aggregation in freshwater mussels, A. plicata and Quadrula spp...... 10 Abstract 10 2-1 Introduction 11 Unique Aspects of Bivalve Physiology 12 2-2 Materials and Methods 17 Study Design 17 Animals 17 Hemolymph Sampling and Processing 18 Treatment Compounds 19 2-3 Results 20 Study One - Temperature 20 Study Two - L-cysteine reconstituted at pH 6.5, 7.0, and 8.0 20 Study Three - L-cysteine exposure up to 24 hours 21 Study Four - Formalin 21 Study Five - Cytochalasin and Study Six - Genistein 22 2-4 Discussion 22 Study One - Temperature 22 Studies Two and Three - L-cysteine 23 Study Four - Formalin 24 Study Five - Cytochalasin B and Study Six - Genistein 24 2-5 Conclusions 26 Chapter 2 Tables and Figures 28 References 37

Chapter 3: Biochemical parameters of freshwater mussel hemolymph, A. plicata and Quadrula spp...... 40 Abstract 40 3-1 Introduction 41 3-2 Materials and Methods 45 Study Design 45 Specimen Collection and Processing 46 Husbandry 47 Statistical Analysis 48 3-3 Results 49 Reference Ranges 49 Sodium and Chloride 49 Potassium and Magnesium 50 Calcium and Phosphorus 51 Glucose 51 Isoenzymes 52 General Health 53 Losses 53 3-4 Discussion 54 3-5 Conclusion 59 Chapter 3 Tables and Figures 60 References 69 Chapter 4: Hematological assessment of freshwater mussels, A. plicata and Quadrula spp...... 71 Abstract 71 4-1 Introduction 72 4-2 Materials and Methods 75 Study Design 75 Specimen Collection and Processing 76 Husbandry 77 Statistical Analysis 78 4-3 Results 79 Reference Ranges 79 Hemocyte Differential 79 Total Hemocyte Count 81 General Health 81 Losses 82 4-4 Discussion 82 Cellular Differentials 82 Total Hemocyte Counts 84 Losses 86 4-5 Conclusions 87 Chapter 4 Tables and Figures 88 References 93

Chapter 5: Conclusion...... 96 5-1 Practical guidelines on how to transport and process hemolymph in order to preserve cellular integrity 96 5-2 Chemical and hematological baseline reference ranges for freshwater mussels 97 5-3 Trends in chemical and cytological parameters in freshwater mussels housed in a captive flow through facility over one year 97 References ...... 99

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List of Tables

Table 2.1: Study One Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes transported at ambient temperature and on ice at 1, 4 and 6 hours. n=3, Quadrula spp...... 28

Table 2.6: Study Six Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes exposed to genistein (35uM or 10µg/ml) for one and three hours. n=5, Amblema plicata...... 30

Table 3.1: Hemocyte biochemical parameters and initial capture measurements from Amblema plicata mussels at baseline, n=21. The first quartile shows that 25% of the data are less than or equal to this value. The third quartile shows that 75% of the data are less than or equal to this value...... 60

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Table 3.3: Electrolytes: Sodium, chloride, potassium, and magnesium levels in Quadrula spp. and Amblema plicata at each time point. Number of each genus sampled at each time point: July A. plicata =21 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4 wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9...... 61

Table 4.3: Percentage of eosinophilic granulocytes, basophilic granulocytes, large agranulocytes, and small agranulocytes represented in Quadrula spp. and Amblema plicata .Number of each genus sampled at each time point: July A. plicata=22 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=14, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=10. .. 89

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List of Figures

Figure 2.1: Study One Results: Effect of temperature on untreated hemocytes over time including: (a) percentage of viable cells, and (b) total hemocyte count of hemolymph stored at ambient temperature(black bars) versus hemolymph stored on ice (grey bars). Means are reported with standard error, medians are represented by solid dot. N=3, Quadrula spp...... 31

Figure 2.2: Study Two Results: Effect of L-cysteine reconstituted at varying levels of pH on mussel hemocytes including: (a) cell viability, and (b) total hemocyte count at pH 6.5 (black bars) pH 7.0 (grey bars) pH 8.0 (dark grey-crosshatched). Means are reported with standard error, medians are represented by solid dot. N=3, Quadrula spp...... 32

Figure 2.3: Study Three Results: Prolonged exposure to L-cysteine up to 24 hrs.: (a) percentage of viability, and (b) total hemocyte count, of L-cysteine treated hemolymph (light grey bars) verses untreated hemolymph (black bars). Means are reported with standard error, medians are represented by a solid dot. N=3, Quadrula spp...... 33

Figure 2.4: Study Four Results: Effect of formalin at one hour post collection including: (a) percentage of viability, and (b) total hemocyte count, of formalin at a 1:1 ratio (50mg/ml) (black bars) and formalin at 1:2.5 (28.5mg/ml) (light grey bars) untreated hemolymph (dark grey bars). Means are reported with standard error brackets, medians are represented by a solid dot. N=5, Amblema plicata...... 34

Figure 2.5: Study Five Results: Effect of Cytochalasin B (1µM or 0.5mg/ml) (1 at one hour and 3 hours post collection including: (a) percentage of viability, and (b) total hemocyte count, of cytochalasin B (black bars) and untreated hemolymph (light grey bars). Means are reported with standard error brackets, medians are represented by a solid dot. N=5, Amblema plicata...... 35

Figure 2.6: Study Six Results: Effect of genistein (35µM or 10mg/ml) at one hour and 3 hours post collection including: (a) percentage of viability, and (b) total hemocyte count, of genistein (black bars) and untreated hemolymph (light grey bars). Means are reported with standard error brackets, medians are represented by a solid dot. N=5, Amblema plicata...... 36

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Figure 3.1: Sodium, chloride, potassium, and magnesium levels in Quadrula spp. and Amblema plicata over time in captivity. Asterisks represent statistically significant differences between genera at that time point (p<0.05). Medians reported by a solid dot. Means and standard errors in brackets reported Black bars marked with hatch mark represent baseline values taken from the animals in the wild. Number of each genus sampled at each time point: July A. plicata =21 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4 wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9...... 65

Figure 3.2: Calcium and phosphorus levels in Quadrula spp. and Amblema plicata over time in captivity. Asterisks represent statistically significant differences between genera at that time point (p<0.05). Medians reported by a solid dot. Means and standard errors reported Black bars marked with hatch mark represent baseline values taken from the animals in the wild. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9...... 66

Figure 3.3: Changes in glucose over time in captivity in Quadrula spp. and Amblema plicata. Medians reported by a solid dot. Means and standard errors in brackets reported Black bars marked with hatch marks represent baseline values taken from the animals in the wild. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9 ...... 67

Figure 3.4: Isoenzyme levels in Quadrula spp. and Amblema plicata over time in captivity. Asterisks represent statistically significant differences between genera at that time point (p <0.05). Medians reported by a solid dot. Means and standard errors reported Black bars marked with hatch mark represent baseline values taken from the animals in the wild. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4 wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9 ...... 68

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Figure 4.1: Eosinophilic granulocytes, basophilic granulocytes, large agranulocytes, and small agranulocytes in Quadrula spp. and Amblema plicata. Baseline Total cell counts per ml of hemolymph for each genus is listed for reference: Quadrula spp.:94,000 cell/ml mean, 65,000 cells/ml median. Amblema plicata: 53,273 cells/ml mean, 36,000 cells/ml median. Asterisks represent statistically significant differences between genera at that time point (p<0.05) except basophils in June (p- value <0.10). Medians reported by a solid dot. Means and standard error brackets reported. Black bars marked with hatch marks represent baseline values taken from the animals in the wild. Note: the scale for small agranulocytes is 10%. Number of each genus sampled at each time point: July A. plicata=22 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=14, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=10...... 91

Figure 4.2: Total cell counts in Quadrula spp. and Amblema plicata over time in captivity. Asterisks represent statistically significant differences between genera at that time point (p<0.10). Medians reported by a solid dot. Means and standard error brackets reported Black bars marked with hatch mark represent baseline values taken from the animals in the wild. Number of each genus sampled at each time point: July A. plicata=22 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=14, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=10...... 92

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1 Chapter 1: Introduction

1-1 Introduction North American freshwater bivalves of the family Unionidae represent one of the most endangered faunas of the world. The Nature Conservancy estimates that 55% of freshwater mussels in North America are extinct or imperiled compared to 7% of the continent’s mammals and birds (Master 1990). In response, a national strategy for the

Conservation of Native Freshwater Mussels was developed in 1998 to provide a framework to prevent further extinction and decline (National Native Mussel

Conservation Committee). This document identified a lack of knowledge of basic biology of freshwater mussels and a need for sensitive health monitoring protocols for wild and captive populations. The development of nonlethal methods of hemolymph collection from mussels has facilitated progress in identifying cytological and chemical indicators of health (Gustafson et al. 2005a, Gustafson et al. 2005b, Burkhard et al.

2009). As habitat pressures continue to increase the survival of many species will come to depend either temporarily or permanently on captivity. Overall, captive management has the unique ability to significantly augment conservation efforts through refugia, intensive propagation, and research. Because components of mussel health have not previously been identified, it is unknown why a high proportion of these normally long- lived animals die within the first year of translocation (Cope et al. 2003). Consequently, more sensitive health surveillance methods are needed to detect declining vitality in

freshwater mussel populations specifically captive populations before fatal end points are reached.

This thesis is an effort to advance knowledge of optimal methods for specimen handling and to develop baseline biochemical and hematological reference ranges from a cohort of freshwater mussels in the wild and follow trends in their biochemistry and hematology for one year post translocation into captivity. The specific objectives of my thesis research were: 1) to determine proper hemolymph handling protocols to preserve cellular integrity and function over time and during transport; 2) to develop baseline biochemical and hematologic reference ranges in a population of wild mussels; and 3) to describe population trends in biochemical and hematological parameters over one year following translocation into captivity.

This chapter provides an introduction to unionids and the genera included in these studies, a background on their decline, and current research needs. Chapter two describes the pertinent gaps in knowledge regarding hemocyte physiology and responses to basic influence (such as temperature and pH), and also describes methods to decrease cellular aggregation using compounds commonly used in leukocyte and platelet research.

Chapter 3 presented baseline reference ranges for 10 biochemical parameters in a cohort of wild freshwater mussels and the results from monitoring fluctuations in hemolymph chemistries over one year in captivity. Chapter 4 reports baseline reference ranges for 4 hemocyte cell types from animals in the wild and shifts in cell types from the population through the first year in captivity.

Introduction to Unionids Two species within the genus Quadrula (Quadrula pustulosa and Quadrula quadrula) and one species within the genus Amblema (Amblema plicata plicata) are represented in this study under the family Unionidae. These species are chosen for this study because they are relatively common in Ohio River systems and have stable population numbers. North America contains the majority of the biodiversity of freshwater mussels, also called naiads, unionids, or clams, with over 281 species and 16 subspecies represented (Williams 1993). Freshwater mussels comprise two major families, Margaritiferidae and Unionidae, which are worldwide in their distribution.

Unionidae is the largest and most diverse family which contains 49 separate genera.

The unionid bivalve genus Quadrula contains three species, Quadrula quadrula,

Quadrula metanerva and Quadrula pustulosa, and 20 recognized subspecies based on shell shape (Simpson 1900; Turgeon et al. 1998). Quadrula quadrula and Quadrula pustulosa are considered by Williams et al. (1993) and Lydeard et al. (1999) to be currently stable. Both species are widespread and relatively common throughout the

Mississippi and Ohio River basin with low genetic distances between populations (Berg

1998). Five species within this genus are federally listed as endangered, five non listed species are considered imperiled, and three species are presumed extinct (Serb et al.

2003). Quadrula quadrula, common name Mapleleaf, has a fairly thick shell that appears square in outline with 2 rows of pustules and a ridged sulcus. Quadrula pustulosa, common name Pimpleback, has a rounded shell usually covered in pustules with a deep open shell cavity. Both species are commonly buried within the substrate and actively move within the sediment. 3

The unionid bivalve genus Amblema contains two species groups, Amblema neislerii, that is listed as endangered and Amblema plicata which contains two species,

Amblema plicata perplicata and Amblema plicata plicata that is currently stable

(Williams 1993). Amblema spp. are larger, more sedentary mussels with three characteristic ridges which earned it its common name, Three ridge. Amblema spp. sit higher in the sediment with their shells exposed to sunlight, encouraging algae growth.

Its ridges are thought to streamline the mussel during times of high flow but also provide a more protected surface area for benthic organisms (Watters 1994).

Biological activities of mussels include filter feeding of phytoplankton and fine particulate organic matter, nutrient cycling of materials into usable forms for other invertebrate species (Spooner 2006). Bivalves are also powerful bioturbators which positively alter the physical structure and the chemical nature of the sediment by releasing nutrients from the sediment into the water column and increasing sediment and water oxygen content (Vaughn et al. 2004). Active mussels and spent shells also provide stabilization of the sediment which increases retention of organic matter (Strayer 2004).

Such biological activities combined with the physical habitat provided by the shells make bivalves important contributors to the general aquatic and benthic community (Spooner

2006). In contrast, epifaunal invasive mussel species such as zebra mussels (Dressinia polymorpha) functionally impact streams and lakes via high rates of filter feeding

(Mellina et al. 1995, MacIsaac 1996). However, they do not support other benthic organisms especially when compared to a biomass of healthy unionid mussels that can

exceed the biomass of all other benthic organisms by an order of magnitude (Layzer et al.

1993).

Threats to freshwater mussel populations are multi-factorial and overwhelming for native populations. Alarming trends in urban and agricultural land use practices, increases in municipal and industrial toxic contaminants, and the rapid spread of deadly invasive species make the development of alternative methods of preservation critical to species survival. Unionid populations are long lived slow growing animals with some species documented to live upwards of 100 years (Strayer et al. 2004). Such life characteristics means that populations are slower to recover from die-off events, over harvest, or invasive competition (Brogan 1993), but also that threats to the population may be more difficult to identify. Dramatic acute losses can be detected with sacrifice of a small percentage of the population for histopathology. However, when attempting to detect long term diffuse effects on population health, prohibitively large sample numbers are needed. Development of sensitive, non-lethal measures of health is needed for epidemiological assessment of captive and wild populations.

Captive facilities are currently called on to provide temporary refugia and assistance in reproduction and research in alternative host fish species. The relocation of freshwater mussels has been used for decades by state and federal agencies as a powerful management tool when critical habitats are threatened (Sheehan et al.1989, Layzer and

Gordon 1993). However, prior to 1995, relocation rates of survival and recovery were as low as 43% (Cope et al.2003). Currently, the survival rate post relocation has improved

to 90% (Cope et al.2003) providing hope that populations can be successfully moved into captivity without prohibitive losses.

Unfortunately there is a dearth of information about the physiology and key components of health in these species and research methods to investigate them are in the early stages of development. The ability of health assessment protocols to be successfully applied to populations depends on the development of reference ranges and accurate identification of the next most crucial research question. This thesis aims to move the knowledge of hematology, clinical pathology, and basic handling protocols for freshwater mussel hemolymph at least a small step forward.

References

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Master, L. 1990.The Imperiled status of North American aquatic animals. Biodiversity Network News. 3: 1-8.

Mellina, E., J.B. Rasmussen, E.L. Mills. 1995. Impact of zebra mussel (Dressinia polymorpha) on phosphorous cycling and chlorophyll in lakes. Canadian Journal of Fisheries and Aquatic Sciences. 52: 2553-2573.

National Native Mussel Conservation Committee 1998. National strategy for the conservation of native freshwater mussels. Journal of Shellfish Research. 17(5): 1419- 1428.

Neves, R.J., A.E. Brogan, J. Williams, S.A. Ahlstedt, P.W. Hartfield. 1997. Status of the aquatic mollusks in the southeastern United States: a downward spiral of diversity. In Benz, G.W. and D.E. Collins (eds.), Aquatic fauna in Peril: The Southeastern Perspective. Special Publication 1, Southeast Aquatic Research Institute: 43-86.

Serb, J.M., J.E. Buhay, C. Lydeard. 2003. Molecular systematics of the North American freshwater bivalve genus Quadrula (Unionidae: Ambleminae) based on mitochondrial ND1 sequences. Molecular Phylogenetics and Evolution. 28(1):1–11.

Simpson, CT. 1900. Synopsis of the naiads or pearly fresh-water mussels. Proc. U.S. Nat Mus. 22: 501-1044.

Spooner, D.E. and C.C. Vaughn. 2006. Context- Dependent Effects of Freshwater Mussels on stream benthic communities. Freshwater Biology. 51: 1016-1024.

Strayer, D.L., J.A. Downing, W.R. Haag, T.L. King, J.B. Layzer, T.J. Newton, S.J. Nichols. 2004. Changing perspectives on pearly mussels, North America’s most imperiled animals. BioScience. 54: 429-439.

Turgeon, D.D., J.F. Quinn Jr., A.E. Bogan, E.V. Coan, F.G. Hochberg, W.G. Lyons, P.M. Mikkelsen, R.J. Neves, C.F.E. Roper, G. Rosenberg, B. Roth, A. Scheltema, F.G. Thompson, M. Vecchinoe, J.D. Williams. 1998. Common and scientific names of aquatic invertebrates from the United States and Canada: mollusks, second ed. American Fisheries Society, Bethesda, MD, Special Publication 26.

Vaughn C.C., K.B. Gido, D.E. Spooner. 2004. Ecosystem processes performed by unionid mussels in stream mesocosms: species roles and effects of abundance. Hydrobiologica. 527:35-47.

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Williams, J.D., M.L.Warren, K.S. Cummings, J.L. Harris, R.J. Neves. 1993. Conservation status of freshwater mussels of the United States and Canada. Fisheries 18(6):22.

2 Chapter 2: Processing methods for hemolymph and efforts to reduce hemocyte aggregation in freshwater mussels, A. plicata and Quadrula spp.

Abstract Hemolymph, the circulatory fluid of bivalves, and the cellular portion called hemocytes, are just beginning to be investigated in freshwater mussels. Recent studies have shown that hemolymph can be safely and repeatedly drawn (Gustafson et al.

2005a). In addition, a schematic for freshwater mussel hemocytes was developed using

L-cysteine as a novel anticoagulant (Burkhard et al. 2009). Due to the marked physiological differences between invertebrate and vertebrate species, handling protocols for hemolymph and hemocyte enumeration cannot be inferred from mammalian based research. This study investigated the optimal processing temperatures for hemolymph, the ideal pH for reconstitution of the anticoagulant L-cysteine, and the effects of prolonged exposure to L-cysteine in-vitro. Towards an optimal method for hemocyte enumeration, cytochalasin B, genistein, and a lower dose of formalin than previously published were evaluated. Total hemocyte count, percentage of viable cells, and presence or absence of cellular debris as an indicator of cell lysis and degranulation were compared for each experiment. Exposure of hemocytes to temperatures below 10 degrees Celsius increased cellular aggregation in-vitro and decreased cellular viability.

Only formalin treatments were found to increase total cell counts. Treatment of hemocytes with formalin, L-cysteine, cytochalasin B, and genistein resulted in a marked production of cellular debris at one hour post exposure. Untreated hemocytes at ambient

temperature can maintain a high level of viability (80-93%) for up to 24 hours.

Therefore, hemocytes should be transported without fixative or anticoagulant at ambient temperature, and it is recommended that hemocytes only be exposed to L-cysteine

(25mg/ml, pH 8.0) just prior to slide preparations for morphological analysis.

2-1 Introduction Methods to assess the health of freshwater mussel bivalves are an emerging research priority as many populations are critically endangered. In mammalian species, a complete blood count is part of a crucial part of a minimum database for evaluation of health, often providing the first sign of abnormalities (Nelson and Couto 2003).

Hemolymph, the circulatory fluid of bivalves, and the cellular portion called hemocytes, are just beginning to be investigated in freshwater mussels. Recent studies have shown that hemolymph can be safely and repeatedly drawn (Gustafson et al. 2005a). In addition, a schematic for freshwater mussel hemocytes was developed using L-cysteine as a novel anticoagulant (Burkhard et al. 2009). As the majority of laboratories are 1-4 hours away from mussel collection sites in the field or captive mussel facilities, identification of methods that will preserve cellular integrity and function are needed prior to conducting further physiological assays.

Though bivalves are considered a primitive life form, preliminary knowledge of molluscan hemocytes have shown remarkable similarity to mammalian cells.

Hemocytes, specifically granulocytes have been shown to be involved in active phagocytosis that can be broken down into recognition, attachment, uptake and killing of

potential pathogens or non-self items (Pipe et al. 1995). Hemocytes are also shown to secrete humoral factors such as lectins (Pipe et al. 1990) and lysozymes (Cheng 1983).

Hemocytes have even been postulated to secrete serine which has protease-like activity associated with virus neutralization (Hughes et al. 1991).

Unique Aspects of Bivalve Physiology Despite these similarities, striking differences exist between mammalian and molluscan physiology that may also affect hemocyte physiology and must be taken into account prior to investigation of hemocyte function and physiology. First, bivalves are ectothermic and heart rate and circulation of hemocytes are sensitive to fluctuating changes in environmental temperature (Fang 1965). Investigators have also noted that lower temperatures directly affect cell adherence to bacteria in-vitro (Foley and Cheng

1975) and are postulated to increase adherence to collection tubes (Burkhard et al. 2009).

Consequently, once hemolymph is removed from the animal it is unknown whether hemolymph should be stored or transported at ambient temperatures or cooled, similar to mammalian cells.

Second, bivalves are sessile creatures that are unable to physically escape potentially harmful changes in their environment and instead they close their valves and maintain homeostasis metabolically (Kohn 1980, Gosling 2003). The signal and mechanism for gape closure is unknown, but bivalves are thought to be highly responsive to changes in pH that may also affect cells in-vitro (Mackie 1989, Gosling 2003, Morton

1970). L-cysteine is an amino acid anticoagulant that comes in a crystalline form and must be re-suspended before use.

Third, cellular enumeration and classification have been significantly challenged by clumping and degranulation in response to fixatives or anticoagulants in marine and freshwater mollusks (Cheng and Foley 1974, Santarem et al.1994, Gustafson 2003). In mammals, coagulation is immediately prevented by drawing blood into tubes with anticoagulants that have an effect on different aspects of platelet function. Mammalian cells can also be exposed to anticoagulant for extended periods at 4° Celsius without an effect on cellular morphologies (Lloyd 1982). However, in bivalves the mechanism for cellular clumping is unknown and treatment with common anticoagulants (sodium citrate and EDTA) and concentrations of L-cysteine >40mg/ml do not reduce clumping and result in hemocyte degranulation and lysis (Burkhard et al. 2009). Mechanical separation by vortexing specimens for 30 seconds was found to partially re-suspend cells, but they would ―settle out‖ and quickly re-aggregate (Burkhard et al. 2009). Mechanical separations plus the addition of L-cysteine at 25mg/ml just prior to preparation of slides was found to significantly reduce cell clumps and maintain cellular integrity for morphological description (Burkhard et al. 2009). The ability to collect hemocytes directly into L-cysteine and keep hemocytes in suspension over time would be greatly valuable for reducing preparation steps. However, the effect of prolonged exposure to L- cysteine is unknown.

Formalin and other formal based fixatives have been used with some success in increasing total cell counts in marine species. However, considerable inter-animal variability in hemocyte enumeration in marine species can still occur as seen in a study examining the effect of copper exposure on total hemocyte counts, in which significant variability was seen in treatment as well as control groups complicating the results (Pipe and Coles 1995). Additional studies show that a common reaction of granulocytes to fixation has been degranulation. Intense degranulation responses have been shown to obscure visualization of hemocytes for enumeration due to an increase in cellular debris as well as to falsely skew enumeration of granulocytes in the total count in Mercinaria mercenaria and Mytilus galloprovincialis (Foley and Cheng 1974, Santarem et al.1994).

In freshwater species, formalin (10%) at a 1:1 ratio to hemolymph has been found to significantly increase cell counts and reduce clumping, however, crenation and degranulation of cells occurred to the extent that performance of a cellular differential was not even attempted (Burkhard et al. 2009). Reducing the concentration of formalin may decrease the effect of fixation on granulocytes and provide a standardized method for enumeration. Consequently, formalin (10% buffered) at a 1:2.5 ratio was used in this study versus a 1:1 ratio used in previous studies to determine if reducing the concentration of formalin may increase cell viability, minimizing the effect on cellular integrity enough for accurate enumeration.

Modern electron microscopy has revealed that meshworks of fine filamentous material exist in a variety of organisms that function as the contractile machinery for cell movement and phagocytosis (Allison et al.1971, Wessells et al. 1971). Pseudopods or

pseudopodia, which are arm-like extensions of the cytoplasm, are thought to play a role in adherence and response to foreign materials, described primarily in granulocytes

(Lopez et al.1997, Cheng and Foley 1975). If pseudopodia are involved in aggregation and have microfilament function, then inhibition of microfilament action may be a successful way to reduce aggregation of cells for more accurate cell counts.

Two compounds, cytochalasin B and genistein, have been shown to affect a variety of cells by altering their chemical or physical structure, primarily through inhibition of microfilament action (Sulahian et al. 2008, Allisson et al. 1971). In hemocytes of the gastropod, Helix pomatia, cytochalasin B was observed to inhibit redistribution of surface molecules by affecting cellular microfilaments (Dageforde et al.

1986) and in Mytilus edulis, pseudopodia inhibition and reduced aggregation of hemocytes was observed after addition of cytochalasin B to cells (Friebel and Renwrantz

1995). Cytochalasin B has been used in human research due to its effect on microfilaments to reduce phagocytosis in macrophages (Axline and Reaven 1974,

Aikawa et al. 1982) and stimulate oxytocin secretion in bovine luteal cells (Shibaya et al.

2005) by reversibly inhibiting microfilament function (Allisson et al. 1971). Genistein is a naturally occurring tyrosine kinase inhibitor and a potent isoflavinoid that has been found to affect phagocytosis by inhibiting scavenger receptors in murine macrophages

(Sulahian et al. 2008) and decreasing platelet aggregation due to an unknown mechanism

(Gottstein et al. 2003). Genistein is also an inhibitor of a transporter mechanism for cellular uptake of glucose in mammalian cells (Vera et al. 1996). In conclusion,

cytochalasin B and/or genistein may be effective in reduction of hemocyte aggregation and provide preliminary knowledge of freshwater mussel cellular physiology.

Most wild and captive mussel collection sites are 1-4 hours from the laboratory.

Consequently, we wanted to know how we should process and transport cells to maintain a high level of viability and minimize aggregation for further physiological testing. The goal of this study was to determine optimal conditions for handling and transport of hemolymph and study the effect of time on hemocytes in-vitro. Several preliminary studies were performed to ask key questions about cellular processing, handling, and physiology: 1) Should cells be collected and transported on ice or at ambient temperature?; 2) What is the optimal pH for reconstitution of L-cysteine?; 3) Does prolonged exposure of hemocytes to L-cysteine affect cellular enumeration and viability?; 4) Can cytochalasin B and/or genistein be used to decrease cell aggregation?;

5) How does treatment with cytochalasin B and genistein compare to fixation with formalin and untreated hemolymph?; 6) Does a lower concentration of formalin decrease the degranulation response seen in hemocytes exposed to formalin at higher doses?; and

7) How long will untreated and treated cells remain viable? All studies measured the total number of live and dead hemocytes over time.

2-2 Materials and Methods

Study Design Hemolymph was obtained for 6 studies. Studies 1 through 3 examined handling of hemolymph during transport including: 1) temperature, 2) pH of reconstituted L- cysteine, 3) prolonged exposure to L-cysteine. Studies 4 through 6 were performed to investigate the following compounds ability to reduce cellular aggregation and their effect on cellular viability including: 4) Formalin at a higher and lower concentration, 5) cytochalasin B, and 6) genistein. An aliquot of untreated hemolymph at ambient temperature was used as a comparison and control for each study. The treatments in each study included: transport of untreated cells on ice or at room temperature, L-cysteine

(25mg/ml) reconstituted at a pH of 6.5, 7.0 and 8.0, and L-cysteine (25mg/ml) exposure up to 24 hours, formalin (50mg/ml), formalin (28.5mg/ml), cytochalasin B (0.5µg/ml), and genistein (10µg/ml). For each treatment and control a total hemocyte count including live and dead cells was performed. Studies 1 through 4, including the temperature, L-cysteine, and formalin studies were performed in the fall of 2008. Study

5 (cytochalasin B) and study 6 (genistein) were performed in the summer of 2009.

Animals Hemolymph was obtained from 10 Amblema plicata and 9 Quadrula including the species Q. quadrula and Q. pustulosa. Studies 1-4, included 3 animals of a mixture of Q. pustulosa or Q. quadrula. Studies 5 and 6 included 5 Amblema plicata each.

Mussels were housed at the Columbus Zoo and Aquarium’s Freshwater Mussel

Conservation and Research Center (FMCRC), located on the Scioto River in Franklin

County, Ohio. General mussel health was evaluated prior to inclusion in the study and weekly for 3 months post experimentation. Animal health was evaluated by observation of burrowing into the substrate, retracting and/or extending the foot, adductor muscle response to mantle touch, and resistance to manual opening of the shell. Mussels included in these experiments were sampled only once.

Hemolymph Sampling and Processing For each study a total of 350µl of hemolymph was drawn with a 1ml syringe with a 25 G 5/8‖ needle from a sinus in the anterior adductor muscle, as previously described

(Gustafson et al. 2005, Burkhard et al. 2009). Samples were transferred to a 1ml microcentrifuge tube and vortexed for 30 seconds immediately after collection. Within

40 minutes of collection, hemolymph was transferred into equal aliquots of 50µL for each treatment at the laboratory. Aliquots of 10µl of hemolymph were stained at a 1:1 ratio with trypan blue exclusion stain (Mp Biomedicals Inc., Solon, OH, USA). The number of live versus dead cells (trypan blue negative and positive, respectively) was quantified using a Neubauer haemocytometer under light microscopy at 1, 4, and 6 hours post collection for all studies except for study 3, which included prolonged exposure to

L-cysteine and had two additional time points at 10 and 24 hours post exposure. The total cell count was calculated based on the dilution factor. Hemocyte viability was reported as a percentage of live cells. Descriptive statistics are used to compare results.

Treatment Compounds L-cysteine (Calbiochem, EMD Biosciences Inc., CA, USA) was reconstituted in nuclease free sterile water to a working solution of 50mg/ml. For study 2, three aliquots of the working concentration were normalized to a pH of 6.5, 7.0 and 8.0. To achieve the final concentration of (25mg/ml), 50µl of hemolymph was added to a tube containing

50µl of L-cysteine (50mg/ml) to obtain a final concentration of L-cysteine at (25mg/ml) at a pH of 6.5, 7.9 and 8.0 respectively. For the high dose of formalin at a 1:1 ratio of formalin to hemolymph, 50 µl of formalin (100mg/ml) (3.7% formaldehyde) (Fisher

Scientific Research, PA, USA; buffered by The Ohio State University Histopathology

Laboratory) were added to 50 µl of hemolymph, for a final concentration of 50mg/ml of formalin (1.85 % formaldehyde). For the low dose of formalin at a 1:2.5 ratio of formalin to hemolymph, 20µl of formalin (100mg/ml) (3.7% formaldehyde) were added to 50µl of hemolymph for a final concentration of 28.5mg/ml formalin (1.0 % formaldehyde). Cytochalasin B (Sigma-Aldrich, MO, USA) was used at a working stock solution of 9.59 mg/ml suspended in DMSO (0.046% DMSO) normalized to a pH of 8.0.

To achieve the final concentration of 0.5mg/ml, 50µl of hemolymph were added to a tube containing 2.63µl of cytochalasin B (9.59mg/ml). Genistein (Sigma-Aldrich, MO, USA) was used at a working concentration of 20mg/ml suspended in DMSO (0.51%) normalized to a pH of 8.0. To obtain the final concentration of genistein at 10mg/ml,

50µl of genistein (20mg/ml) were added to 50µl of hemolymph.

2-3 Results

Study One - Temperature The viability of mussel hemocytes stored at ambient temperature was consistently higher than of those stored on ice throughout the study. Viability of hemocytes stored at ambient temperature was 7% 19%, and 22% higher than hemocytes on ice at 1, 4, and 6 hours respectively (Figure 2.1a). Total cell counts in hemocytes at ambient temperature were ~5 times higher than cells on ice at one hour. At 4 hours, both groups showed marked increases in total cell count, with an increase of 69% in cells at ambient temperature compared to a 150% increase in cells on ice (Figure 2.1b). Cell counts in both groups were steady or increased between 4 hour and 6 hours, but the number of dead cells increased dropping the average percentage of viability from 92% viable cells at 4 hours to 72% at 6 hours at ambient temperature and from 78% viable cells to a mean of

60% in cells on ice at 4 and 6 hours respectively (Table 2.1).

Study Two - L-cysteine reconstituted at pH 6.5, 7.0, and 8.0 The lowest mean viability percentage and lowest mean hemocyte count at all time points was observed in hemocytes reconstituted in L-cysteine at a pH of 6.5 (Figures 2.2 a, b). L-cysteine at a pH of 8.0 maintained the highest viability and average total cell count at one hour (81% and 132,000cells/ml) and at 4 hours (85% and 105,000 cells/ml).

At 6 hours, viability of L-cysteine at pH of 7.0 and total cell count (74% and 102,000 cells/ml) was slightly higher than L-cysteine at a pH of 8.0 (72% and 74,000cells/ml)

(Figures 2.2 a, b). A high amount of cellular debris in L-cysteine at a pH of 6.5 and pH

of 7.0 obscured the ability to enumerate cells at 4 and 6 hours. Hemocytes maintained in

L-cysteine at a pH 8.0 appeared intact with no cellular debris observed at any time point

(Table 2.2).

Study Three - L-cysteine exposure up to 24 hours Untreated cells maintained greater than 90% viability at all time points throughout the study with a mean viability of 96% at one hour and 93% at 24 hours (Figure 2.3a).

Mean viability in cells exposed to L-cysteine were 34% which was 30% lower than untreated cells at 1 and 4 hours. Between 4 and 6 hours, viability of hemocytes exposed to L-cysteine dropped to less than 50% and at 24 hours was less than 40%. Total cell count in L-cysteine treated cells was 38% higher than the average cell count in untreated cells at one hour (Figures 2.3 b). At 4 hours, L-cysteine treated cell count was only 10% higher than untreated cells. An increase in cellular debris was noted at 1, 4, and 6 hours post exposure to L-cysteine (Table 2.3)

Study Four - Formalin Hemocytes exposed to formalin at a 1:1 ratio were 62% viable at one hour post exposure versus 70% in formalin at a 1: 2.5 ratio and 83% in untreated cells. The average total cell count in formalin (1:1) treated cells was 60% higher than untreated cells and 55% higher in formalin (1:2.5) (Figure 2.4 a,b). Severe amounts of cellular debris prohibited enumeration beyond one hour of exposure in both groups (Table 2.4).

Study Five - Cytochalasin and Study Six - Genistein Less than half of the cells treated with genistein were viable at one hour compared to cytochalasin B-treated hemocytes which were 62% viable, compared to 86% viability in untreated cells (Figures 2.5 a, b and Table 2.5). Treatment with cytochalasin B resulted in a 33% decrease in cell count compared to untreated cells at one hour.

Genistein resulted in a 4% increase in cell count compared to untreated cells at one hour

(Figures 2.6 a,b and Table 2.6). Both groups had a moderate amount of cellular debris at one hour, thus cell counts were repeated at 3 hours instead of 4 hours. Genistein and cytochalasin B treated hemocytes showed an increase in the amount of cellular debris at 3 hours, almost precluding enumeration. At 3 hours, total number of cells/ml enumerated for cytochalasin B-treated cells was 60% lower than untreated cells and the total count with genistein was 12% lower than that of untreated cells. At 3 hours, the viability in cytochalasin B cells was 66% compared to genistein at 46% and untreated cells at 80%.

2-4 Discussion

Study One - Temperature The level of cell aggregation appears to decrease with time regardless of transport temperature, as seen at 4 hours when total cell counts increase in both hemocytes stored at ambient temperature and on ice. The percentage of viable cells also increases in both groups between 1 and 4 hours which indicates that while cells may lose their ability to aggregate they are not dead and not up-taking trypan blue stain. Additionally, the results

suggest that untreated hemolymph kept at ambient temperature can maintain a high level of viability (greater than 90%) in-vitro for up to 4 hours.

Studies Two and Three - L-cysteine The L-cysteine pH experiments revealed that reconstitution of 50mg/ml

L-cysteine at a pH of 8.0 added at a 1:1 ratio with hemolymph resulted in the highest cell viability and highest total cell count. In addition, hemolymph treated with L-cysteine at a pH of 8.0 was devoid of cellular debris even at 6 hours post exposure, indicating that cells were likely intact and not stimulated to degranulate. Water from the captive facility routinely ranged from a pH of 8.0 and pH of 8.3. Animals in this study were apparently acclimated to higher pH levels and when exposed to L-cysteine at a pH of 6.5 or 7.0 hemocytes were stimulated to aggregate and degranulate.

The L-cysteine exposure experiment was performed using L-cysteine reconstituted at a pH of 8.0 and transported at ambient temperature based on the results of the previous study. Exposure to L-cysteine (25mg/ml) at a pH of 8.0 for one hour resulted in an average hemocyte viability of around 80% as seen in the pH study.

Untreated hemocytes maintained an average viability of over 90% throughout the study.

At 4 hours total cell count differences between treated and untreated cells were negligible. Cellular response to L-cysteine at one hour varied in L-cysteine at pH of 8.0 in each study. In the pH study, L-cysteine treated cells were observed to be free of cellular debris up to 6 hours, whereas in the length of exposure study L-cysteine at the same concentration and pH showed marked cellular debris one hour post-exposure. The

reason for the variation described above is unknown. The studies were performed at the same time of year less than a week apart from one another; therefore, differences are not explained by season or temperature, but could be related to inter-animal variability, especially given the small number of animals (n=3) included in the study.

Study Four - Formalin Both concentrations of formalin (high at 1:1 or low at 1:2.5) appeared to decrease cellular aggregation, but they also stimulated cells to degranulate. Formal-based fixatives, in theory, should rapidly kill cells and result in roughly less than 10% of viable cells. In this study, formalin-treated cells had greater than 60% viability. This indicated that formalin only destroyed the cellular membrane of roughly 40% of the cells, which allows the uptake of trypan blue stain. Formalin, because it contains formaldehyde, should also kill cells fast enough to not elicit a degranulation response. The higher concentration of formalin destroyed 10% more of the population’s cells when compared to the lower concentration, which indicated that there was a possible dose effect.

Study Five - Cytochalasin B and Study Six - Genistein Exposure to genistein increased total cell counts by less than 5% compared to untreated cells at one hour and exposure to cytochalasin B resulted in a decrease in cell counts compared to controls. Four hours post exposure, cytochalasin B and genistein- treated cells demonstrated lower cell counts than untreated cells and marked degranulation as well as an increase in cellular debris was observed. These results are in

contrast with results from Friebal and Renwrantz (1995) who reported that a strong reduction in hemocyte aggregation was observed after the addition of cytochalasin B, purportedly due to inhibition of pseudopodia activity in Mytilus edulis hemocytes. The reason for these differential findings is unknown. It may be due to general differences between marine and freshwater species or it may be due to differences in the role of pseudopodia in marine versus freshwater hemocytes. For example, pseudopodia in freshwater mussels may play a role in phagocytosis but not cellular adherence. Whether phagocytosis was inhibited cannot be determined under light microscopy and would require additional study under electron microscopy.

Additionally, this project was not designed to study whether freshwater hemocytes have microfilaments or not. Microfilaments may be present but involved in other hemocyte processes such as lysosomal release. For example, it was found in mammalian cells that when challenged with zymosan particles, cytochalasin B blocks neutrophils from phagocytizing and enhances the release of lysosomal enzymes from neutrophils. The mechanism is unknown but is thought to be due to an effect on microfilaments that are responsible for translocation of lysosomes (Zurier et al. 1973).

Future studies are needed to determine if lysosomal release is increased in response to cytochalasin B exposure, which may provide a case for the presence of microtubules.

Overall, there is a positive association of increased cell counts with time in untreated hemolymph, which potentially indicates that cell aggregation could be dependent on an energy source that is depleted over time. Untreated hemolymph is capable of maintaining >85% viability for up to 24 hours.

2-5 Conclusions This is the first study to investigate processing variables that may impact freshwater mussel hemocyte cell counts and cell viability after collection and during transport. Hemocytes should be transported without anticoagulant at ambient temperature. Any diluents or additives to hemolymph need to be at a pH most similar to physiological levels. Transporting hemolymph on ice appears to increase hemocyte aggregation in-vitro, thereby decreasing total cell counts. Total cell count also increased over time, indicating that cellular aggregation may be dependent on an energy source that is depleted over time. Cell viability decreased after 4 hours regardless of whether the cells were treated or untreated, indicating that future studies that may look at cell physiology and function need to be analyzed within 4 hours after collection. An increase in degranulation can occur after exposure to L-cysteine for one hour and a reduction in the number of viable cells by ~20% can be seen. Consequently, it is recommended that freshwater hemocytes be transported untreated at ambient temperature and that L- cysteine (25mg/ml) at pH of 8.0 be added just prior to preparation of slides for cytocentrifugation to minimize effects on cellular morphology.

Cytochalasin B and genistein appeared to increase cellular debris and degranulation despite being reconstituted at a pH of 8.0 and had a negative effect on total cell count. Further studies are needed to confirm the presence or absence of microfilaments and determine their action in freshwater bivalves. Additional studies are needed to identify the mechanisms of cellular aggregation. Cellular aggregation is a

barrier to more standardized enumeration, but currently counting cells using untreated hemolymph 1 hour post collection and thoroughly vortexing cells to mechanically release cells into suspension prior to counting is the best method to date.

Chapter 2 Tables and Figures

Table 2.1: Study One Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes transported at ambient temperature and on ice at 1, 4 and 6 hours. n=3, Quadrula spp. Study 1: Temperature % viable cells/ml Time (hrs) Ambient Ice Ambient Ice Mean 76.7 71.7 34,667 16,667 1 Median 80.0 90.0 40,000 8,000 Stdev 25.2 40.7 24,440 20,429 Mean 92.5 77.8 58,667 42,000 4 Median 92.9 83.3 26,000 20,000 Stdev 7.7 25.5 72,728 47,032 Mean 72.2 59.2 52,667 64,667 6 Median 66.7 68.4 22,000 38,000 Stdev 21.0 35.4 67,449 71,815

Table 2.2: Study Two Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes transported at ambient temperature in L-cysteine (25mg/ml) reconstituted at a pH of 6.5, 7.0, and 8.0. Cell counts and percentage of viable cells were measured at 1, 4, and 6 hours. n=3, Quadrula spp. Study 2: L-cysteine pH % viable cells/ml Time (hrs) pH 6.5 7.0 8.0 6.5 7.0 8.0 Mean 47.5 66.9 80.6 57,333 76,000 132,000 1 Median 67.5 58.8 78.7 80,000 88,000 148,000 Stdev 41.3 19.9 8.7 46,361 33,645 65,483 Mean 67.7 77.6 86.7 83,333 84,667 105,333 4 Median 70.0 77.0 86.0 86,000 102,000 98,000 Stdev 4.9 5.0 7.0 54,049 33,546 77,261 Mean 67.0 75.0 73.3 80,667 74,000 102,000 6 Median 71.0 77.0 74.0 68,000 94,000 94,000 Stdev 7.8 7.2 7.0 52,166 36,387 44,542

Table 2.3: Study Three Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes exposed to L-cysteine (25mg/ml, pH 8.0) for 1, 4, 6, 10 and 24 hours. n=3, Quadrula spp. Study 3: L-cysteine exposure % viable cells/ml Time (hrs) Untreated L-cysteine Untreated L-cysteine Mean 96.4 63.2 92,667 123,333 1 Median 96.2 62.4 106,000 96,000 Stdev 3.53 11.48 84,790 69,176 Mean 91.8 62.8 135,333 148,000 4 Median 93.6 53.4 156,000 188,000 Stdev 9.18 18.4 108,487 85,346 Mean 92.8 48.9 141,333 152,000 6 Median 93.8 50.9 148,000 136,000 Stdev 2.4 6.9 106,157 77,253 Mean 93.1 43.4 151,333 103,333 10 Median 95.6 41.2 144,000 102,000 Stdev 8.7 4.4 89,226 20,033 Mean 92.8 37.8 234,667 136,000 24 Median 92.8 38.9 220,000 146,000 Stdev 2.8 4.5 214,377 59,632

Table 2.4: Study Four Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes exposed to formalin for one hour at a high concentration (1:1 ratio formalin to hemolymph) and a low concentration (1:2.5 formalin to hemolymph). n=5, Amblema plicata. Study 4: Formalin at 1 hour % viable cells/ml 1:1 1:2.5 Control 1:1 1:2.5 Control Mean 61.7 69.0 82.8 69,600 67,200 43,200 Median 64.7 82.6 88.9 52,000 50,000 42,000 Stdev 16.8 25.8 17.6 43,552 43,165 21,288

Table 2.5: Study Five Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes exposed to cytochalasin B (1uM or 0.5µg/ml) for one and three hours. n=5, Amblema plicata. Study 5: Cytochalasin B % viable cells/ml Time (hrs) 1µM Control 1µM Control Mean 61.9 85.8 48,800 64,800 1 Median 52.3 91.2 42,000 54,000 Stdev 19.7 13.5 13,535 30,646 Mean 66.4 80.0 48,400 77,200 3 Median 60.8 84.3 46,000 92,000 Stdev 18.1 9.8 20,513 25,519

Table 2.6: Study Six Results. Percentage of viable cells and total cell count (cells/ml) for hemocytes exposed to genistein (35uM or 10µg/ml) for one and three hours. n=5, Amblema plicata. Study 6: Genistein % viable cells/ml Time (hrs) 35µM Control 35µM Control Mean 40.5 72.8 19,200 18,400 1 Median 41.2 70.8 20,000 18,000 Stdev 11.8 5.9 2,280 5,727 Mean 46.3 60.2 24,000 26,800 3 Median 42.3 53.8 26,000 26,000 Stdev 16.4 19.8 11,314 3,033

(a) Effect of temperature on viability (b) Effect of temperature on hemocyte count

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0 0 Treatment d s d s d s d s d s Treatment d s d s d s d s d s te y te y te y te y t e y te y te y te y te y t e y a c a c a c a c a c a c a c a c a c a c e L e L e L e L e L e L e L e L e L e L tr tr tr tr tr tr tr tr tr tr n n n n n n n n n n u u u u u u u u u u

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(a) Effect of cytochalasin B on viability (b) Effect of cytochalasin B on total hemocyte count

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References Aikawa, M., L.D. Hendricks, Y Ito, and M. Jagusiak. 1982. Interactions between macrophage like cells and Leishmania brazilensis in vitro. American Journal of Pathologists. 108(1):50-59.

Allison, A. C., P. Davies, S. de Petris. 1971. Role of contractile microfilaments in macrophage movement and endocytosis. Nature: New Biol. (London). 232, 153.

Axline, S.G and E.P Reaven. 1974. Membrane mobility of the cultured macrophage by cytochalasin B: the role of subplasmalemmal microfilaments. The Journal of Cell Biology. 62:647-659.

Bogolyubova, N. A., I. O Bogolyubova, V.N Parfenov, G.G Sekirina. 1999 Peculiarities of structural and functional organization of two-cell mouse embryos exposed to certain inhibitors of cell proliferation. Tsitologiya. 41(8): 698-706.

Cheng, T. C. 1983. The role of lysosomes in molluscan inflammation. Amer. Zool. 23: 129-144.

Cheng, T.C. and D.A. Foley. 1975. Hemolymph Cells of the Bivalve Mollusc Mercenaria mercenaria: An Electron Microscopical Study. Journal of Invertebrate Pathology. 26: 341-351.

Dageforde, S., A. Schmucker, L. Renwrantz. 1986. Capping of cell surface receptors on blood cells from the molluscs Helix pomatia (Gastropoda) and Mytilus edulis (Lamellibranchiata). Eur. J. Cell Biol. 41:113-120.

Feng, S.Y. 1965. Heart rate and leukocyte circulation in Crassostrea virginica. Biological Bulletin. 128:198-210.

Foley, D.A. and T.C Cheng. 1974. Morphology, hematologic parameters, and behavior of hemolymph cells of the Quahog clam, Mercenaria mercenaria. Biological Bulletin. 146:343-356.

Foley, D.A. and T.C. Cheng. 1975. A quantitative study of phagocytosis by hemolymph cells of the pelecypods Crassostrea virginica and Mercenaria mercenaria. Journal of Invertebrate Pathology. 25:189-197.

Friebel, B. and L. Renwrantz.1995.Applicaiton of density gradient centrifugation for separation of eosinophilic and basophilic hemocytes from Mytilus edulis and characterization of both cell groups. Comparative Biochemical Physiology part A. 112(1): 81-90.

Gosling, E. 2003. Bivalve mollusks: Biology, ecology and culture. Blackwell Publishing, Malden MA.

Gottstein, N., B.A. Ewins, C. Eccleston, G.P. Hubbard, I.C. Kavanagh, A.M. Minihane, P.D. Weinberg, G. Rimbac. 2003. Effect of genistein and daidzein on platelet aggregation and monocyte and endothelial function. British Journal of Nutrition. 89:607–615.

Gustafson, L.L. ―Nonlethal Health Assessment of the Freshwater Mussel Elliptio complanata.‖ Diss. North Carolina State U. 2003.

Hughes, T.K., E.M. Smith, J.A. Barnett, R. Charles, G.B. Stefano. 1991. Lps stimulated invertebrate hemocytes: a role for immunoreactive TNF and IL-1. Developmental and Comparative Immunology. 15:117-122.

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Lloyd E. 1982. The deterioration of leukocyte morphology with time: Its effect on the differential count. Lab Perspect 1(1):13-16

Lopez, C., M.J. Caballal, C. Azevedo, A. Villalba. 1997. Morphological characterization of the hemocytes of the clam, Ruditapes decussates (Mollusca: Bivalvia). Journal of Invertebrate Pathology. 69:51-57.

Mackie, G.L. 1989. Tolerances of Five Benthic Invertebrates to Hydrogen Ions and Metals (Cd, Pb, AI). Archives of Environmental Contamination and Toxicology. 18: 215- 223

Morton, B. 1970. The tidal rhythm and rhythm of feeding and digestion in Cardium edule. Journal of the Marine Biological Association of the United Kingdom. 50:499-512.

National Native Mussel Conservation Committee 1998. National strategy for the conservation of native freshwater mussels. Journal of Shellfish Research. 17(5): 1419- 1428.

Nelson, R.W., C.G. Couto. Small animal internal medicine. 3rd ed. St. Louis: Mosby, 2003. 1362p.

Neves, R.J. 2010. Partnerships for Ohio River Mussels. (Online) United States Fish and Wildlife Service. http://www.fws.gov/midwest/endangered/clams/ohio_rvr.html accessed November 11, 2010.

Pipe, R.K. 1990. Differential binding of lectins to haemocytes of the mussel Mytilus edulis. Cell Tissue Res. 261:261–268.

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Pipe, R.K., J.A. Coles, M.E. Thomas, V.U. Fossato, and A.L. Pulsford. 1995. Evidence for environmental derived immunomodulation in mussels from the Venice lagoon. Aquatic Toxicology. 32: 59-73.

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3 Chapter 3: Biochemical parameters of freshwater mussel hemolymph, A. plicata and Quadrula spp.

Abstract Biodiversity within freshwater ecosystems has suffered due to widespread reduction in water and habitat quality due to detrimental land-use practices, damming and channelization of major river systems, and pollution. Freshwater mussels have lost almost 75% of the biodiversity that was present in the early 1900s. Improved health surveillance of remaining populations is needed to recognize a decrease in vitality before further losses occur. Hemolymph chemistry may be a useful non-lethal indicator of bivalve physiological processes once reference ranges are established to differentiate normal from abnormal fluctuations. The purpose of this study was to establish reference ranges for the following measurable components in hemolymph: sodium, chloride, magnesium, phosphorus, potassium, calcium, glucose, and isoenzymes alanine aminotransferase (ALT), aspartate aminotransferase (AST), and alkaline phosphatase

(ALP) in a population of mussels in the wild and follow changes in hemolymph from the same population of freshwater mussels translocated into captivity over one year.

Hemolymph from forty animals of three species, Amblema plicata, Quadrula quadrula, and Quadrula pustulosa was collected in July of 2008 from the Muskingum

River in Devola Ohio to develop baseline reference ranges. Thirty-nine of those forty animals were translocated into captivity, with nine animals serving as captive controls without any sampling. Animals were sampled biweekly for the first month and then

quarterly over one year. Significant differences in sodium, potassium, chloride, magnesium, ALT, AST, and ALP were found between genera, A. plicata and Quadrula spp. at baseline (p <0.05). Both genera showed declines in sodium, chloride, and potassium in the first month in captivity and had marked increases in all electrolyte values in November, five months after transport to captivity. Calcium and glucose values remained steady in the population until the last collection point in June when values in both parameters declined. Phosphorus levels increased in both genera with significantly higher levels seen in A. plicata in February (p <0.05). Death losses occurred during the study in greater proportion after winter sampling time points, potentially indicating that sampling during winter quiescence is not recommended. This study has built the foundation for development of broader reference ranges for these species.

3-1 Introduction The proportion of wildlife species at risk is disproportionately large among aquatic species compared to terrestrial species. The taxon at the top of the list is North

America’s freshwater mussels with almost 70% of species presumed extinct, imperiled, or vulnerable (Williams et al.1990). Their conservation status represents an immediate need for sensitive measures of declining vitality. In wild populations, evidence of morbidity or declining vitality in populations can be subtle and nonspecific and may not manifest in clinical disease, thus requiring large sample sizes to identify key threats to population health. In mammals and other vertebrates, tracking the level and trends of chemical parameters within blood can be reflective of fluctuations in health and represent

a safe repeatable method for epidemiologic studies. In order to differentiate normal from abnormal fluctuations, reliable chemical reference ranges are needed. Once developed, chemical parameters in hemolymph could be part of a minimum diagnostic database for an effective health assessment protocol for wild and captive populations.

Establishment of reference ranges have been attempted in wild free ranging freshwater bivalves but have been complicated by dynamic environmental fluctuations in water quality (Gustafson et al.2005, Wilbur 1993). Bivalves are sessile and ectothermic creatures that are highly susceptible to changes in their surrounding habitat (Dietz 1974).

Bivalves maintain homeostasis by metabolic compensation compared to other aquatic species that can move within the water column to evade suboptimal conditions.

Consequently, basal metabolic rates, gas and ion exchanges, filtration rate, feeding behaviors, and reproductive activities of freshwater mussels are influenced by ambient temperatures and water quality (Gustafson Dissertation 2003). In order to develop a functional and powerful health assessment protocol for freshwater mussels, special consideration needs to be paid to seasonal, reproductive, and environmental variability requiring frequent sampling. Animals kept in artificial laboratory environments have highly controlled environmental conditions but the results have potentially limited application back to the populations of interest (Martem'yanov 2000). Most field-based studies are conducted only during warmer months due to favorable conditions and temperatures for workers, missing valuable life data and potentially creating an incomplete picture of overall health (Gustafson Dissertation 2003). Development of reference ranges in a flow-through captive facility could serve to keep animals in an

environment where they are still exposed to a natural water source, but wide fluctuations in environmental parameters are attenuated and year round access for sampling is secured.

Evaluating the chemical components in circulation in mammalian species is a critical part of routine health assessment and is the first step to detection of alterations in health when clinical signs appear. For veterinarians and human physicians, the diagnostic process combines a through history, a complete physical exam, and baseline laboratory tests to assess general immune system and organ function. Freshwater mussel clinical signs are often very subtle and thorough physical exam is more challenging in molluscan species encased in a hard shell. Physical examination is therefore limited to subjective evaluation of placement within the substrate, foot turgidity, and adductor response. Methods to assess health have been limited to invasive or lethal techniques, requiring sacrifice of a representative portion of the population for histopathology.

Development of a chemistry panel for freshwater mussels may be an effective non- invasive tool that can be used to monitor normal physiological processes and diagnose abnormalities. Previous studies in freshwater bivalves that have evaluated tissue, mantle cavity fluid, and hemolymph for chemical constituents have found that calcium, glucose, sodium, potassium, and magnesium values shift in response to disease states (Pekkarinen 1997) and stress factors, such as translocation (Comibra et al.1993,

Martem’yanov 2000) or anoxic conditions (Wilbur 1993, Dietz 1974). Handling and transportation are thought to disturb normal values of sodium, potassium, calcium and glucose (Pekkarinen & Suoranta 1995) for up to 18 days post translocation (Dietz 1979).

In addition to calcium, magnesium, and glucose, freshwater mussel hemolymph has also been shown to have detectable levels of phosphorus, total protein, and aspartate aminotransferase (Gustafson et al.2005b).

In order to move closer to development of a sensitive non -lethal method for health surveillance, this study aimed to 1) provide initial reference ranges for chemistry values from two common genera of freshwater mussels collected in the wild; and 2) develop an understanding of chemical fluctuations over one year through the use of a captive flow through facility. Once translocated into captivity after baseline sampling in the wild, individuals were allowed to acclimate for 14 days then hemolymph samples were drawn in captivity in Early August, Late August, November, February and June which corresponds to 2 weeks, 1 month, 5 months, 8 months and 11 months post translocation, respectively. Chemical constituents measured in hemolymph and analyzed at each time point included: sodium, chloride, potassium, phosphorus, magnesium, calcium, glucose, alanine aminotransferase (ALT), aspartate aminotransferase (AST), and alkaline phosphatase (ALP).

3-2 Materials and Methods

Study Design Forty wild mussels from two genera, Quadrula and Amblema, and three species,

Q. quadrula, Q. pustulosa, and A. plicata, were collected on July 23, 2008 at Muskingum

River, Devola, OH. Thirty nine of these subjects were successfully sampled and included in the baseline data group; an inadequate amount of hemolymph was obtained from one subject.

Thirty subjects of the original thirty-nine were transported into captivity for repeated sampling. The other nine animals were transported into captivity for observation but were not sampled or handled for the duration of the study. The captive control group consisted of 5 A. plicata, 3 Q. quadrula, and 1 Q. pustulosa. The captive sampling group was comprised of 16 A. plicata, 10 Q. quadrula, and 4 Q. pustulosa.

Animals were allowed to acclimate for 14 days prior to further sampling and were sampled in captivity in early August, late August, November, February, and June. At each time point the other nine animals were observed for a general health exam only.

Mussel health was evaluated by observing an animal’s ability to maintain their position within the substrate and retract and/or extend the foot, as well as adductor muscle response to mantle cavity touch and resistance to manual opening of the shell. Any animal that was unable to exhibit the characteristics above for more than 12-24 hours were presumed dead and removed from the collection and placed into fixative (10% buffered formalin, Fisher Scientific, USA). Animals that had tears in the adductor muscle were not sampled at that time point to minimize further damage and due to concerns that the sinus and the resultant sample may be diluted by mantel cavity fluid. 45

Specimen Collection and Processing Approximately 900µl of hemolymph were collected from each mussel at each time point. Hemolymph was drawn with a 1ml syringe with a 25 G 5/8‖ needle from a sinus in the anterior adductor muscle, as previously described (Gustafson et al. 2005;

Burkhard et al.2009). During the initial collection, mussels were weighed to the nearest

0.1g (Ohaus SP 202 Scout Pro, Ohaus, USA) and shell dimensions (length, width, height) were measured to the nearest mm with vernier calipers (Dial 0-6 inch Calipers,

Craftsman, USA). Shell length is a common measurement used to compare molluscan species (Molina et al.2001). Individual animals were marked with a uniquely numbered vinyl identification tag (Floy tag, USA) secured to the shell non-invasively with underwater epoxy (H2Hold, Devcon, USA). Animals were removed from the substrate with a maximum of 3 minutes of emersion time during sampling and tagging.

Hemolymph samples were processed within 45-60 minutes after collection and kept at ambient temperature throughout processing. To remove the cellular fraction from the supernatant, samples were individually vortexed for 30 seconds and centrifuged for 5 min at 1000 x g. The supernatant was then pipetted into cryovials (Fisher Scientific) and placed on dry ice until stored in a -20ºC freezer. At the end of the year all frozen supernatant samples were thawed in groups of 10 and analyzed on an automated clinical analyzer (Hitachi 911, Roche, USA). The following chemical constituents of freshwater mussel hemolymph were analyzed: chloride, phosphorus, sodium, potassium, calcium, magnesium, glucose; alanine aminotransferase (ALT), aspartate aminotransferase (AST), and alkaline phosphatase (ALP).

Husbandry All animals were housed at the Columbus Zoo and Aquarium’s Freshwater

Mussel Conservation and Research Center (FMCRC). Water quality parameters and mortalities were recorded weekly. All mussels were kept in 4 x 3 x 2ft tanks with 8-12 inches of sand and gravel for substrate. The system contains a constant total water volume of ~10,000 gallons with a water surface area of 495 sq. ft. The number of animals housed in the facility ranges from 1000-2000, depending on the time of year.

The facility is a flow through system that utilizes water from the Scioto River pumped through a large 6 ft tower with biological filtration. The facility has the ability to change to a re-circulating system if needed. The average pH of the water during this study ranged from 7.5-8.3 with an osmolality of 25mmol/kg. The pH and hardness varies with natural fluctuations in the river, and is not modified by the system. Ammonia in the

Scioto River ranges from 0.001 to 0.8 ppm depending on rain events and run-off from agricultural lands up river. During environmental events that could compromise the water quality (ammonia concentration or known pollutants) the river water is re- circulated within the initial collection tower up to 3 times to effectively reduce ammonia to tolerable levels between 0.0 – 0.05ppm before contact with the animal collection. The average water and ambient temperature in the facility was 17ºC with lows of 6 ºC recorded over winter. Dissolved oxygen was consistently above 95% saturation throughout the study.

Statistical Analysis General descriptive trends for each biochemical parameter at baseline and over one year are presented. Genera differences appeared to exist in biochemical parameter levels in the baseline data, which was an unexpected result. To determine if these differences were statistically significant, the Mann-Whitney test was used to evaluate differences in biochemical parameter medians between Quadrula spp and Amblema plicata at baseline and each subsequent time point, a p-value <0.05 was considered significant (Fagerland and Sandvik 2009). Quadrula pustulosa and Quadrula quadrula were pooled for comparisons against Amblema due to the low numbers of Quadrula pustulosa (n<5) included in the study. An association between adductor muscle pathology (including both tears and needle tracks) and death as an outcome was evaluated using a Chi square measure of association test with a Yates correction. Initial and final size measurements of both the control group and the sample group were analyzed for growth using paired t- tests. Statistical analysis was performed using

Minitab 16 Statistical Software (State College, PA: Minitab, Inc. USA).

3-3 Results

Reference Ranges Reference ranges for biochemical parameters in freshwater mussel hemolymph are presented for each genus, due to statistical differences between genera that were found unexpectedly at baseline for the following ions: sodium, potassium, chloride, and magnesium and for all three isoenzymes: ALT, AST, and ALP (p-value<0.05). Animal weights and shell lengths are included to describe the reference populations (Tables 3.1 and 3.2).

Sodium and Chloride Both genera show a marked decline in mean sodium and chloride values after the first month of captivity with a more marked decline demonstrated in A. plicata. After the first 14 days in captivity A. plicata mean sodium values decreased by 42.7% and mean chloride values decreased by 46.7%. In Quadrula spp. a 21.5% decrease in mean sodium values and 27.4% decline in mean chloride values were observed. In Quadrula spp. mean sodium and chloride values in hemolymph appear to have recovered to levels similar to baseline values at some point between 1-5 months in captivity, and even exceed mean baseline levels by 20.2% for sodium and 7% for chloride at 5 months versus baseline means respectively. Quadrula spp. maintained 92% – 97% of their original baseline sodium and chloride values at 8 and 11months post-translocation.

Comparatively, between 1 and 5 months in captivity, Amblema’s population means did rise by 3.24 mmol/L for sodium and by 2.31mmol/L for chloride, but the population means for sodium and chloride were still roughly only 82% and 78% of their respective

baseline values. From November to June means for Amblema plicata declined to approximately 52.6% and 60% of their original baseline values for sodium and chloride

(Table 3.3).

Unexpected genera differences between A. plicata and Quadrula spp. were found at baseline and at each time point. For example, Amblema plicata mussels had higher sodium and chloride values at baseline than Quadrula spp. (sodium p=0.0025, chloride p=0.0036; Figure 3.1). Quadrula spp. have higher values for both sodium and chloride at

5 months (sodium p=0.003, chloride p=0.000) and 11 months (sodium p=0.008, chloride p=0.010) in captivity compared to Amblema.

Potassium and Magnesium Magnesium values in both genera increased post-translocation and peaked at 5 months. From baseline to 5 months post translocation, the mean magnesium values increased by 63% in A. plicata and increased by 65.6% in Quadrula spp. Potassium levels in both genera had an appreciably small measurable volume, as mean values are less than 1mmol/L (Table 3.3).

Unexpectedly, potassium and magnesium concentrations differed at baseline and over time in captivity between the genera (Figure 3.1). A. plicata had higher median potassium values at baseline and in November than Quadrula spp., whereas Quadrula spp. had much higher values of magnesium at baseline, early August, and November than

A. plicata (p value < 0.05).

Calcium and Phosphorus Calcium means appeared to be stable throughout captivity from an initial value of

19.98 mg/dL in July to 20.69 mg/dL in November and 20.95mg/dL in February (Figure

3.2). Calcium values then dropped by 13% from February to June, with the lowest value of 18.21 mg/dL in June after 11 months in captivity. Calcium median levels did not vary significantly by genus at any time point (Table 3.4).

Phosphorus levels in both genera were present at an appreciably small measurable concentration, as mean values are less than 1 mg/dL (Figure 3.2). Phosphorus levels rose post translocation in both genera, however, phosphorus increased over time at a higher percentage in A. plicata than Quadrula spp. From baseline in July to November, phosphorus values increase from 0.43 mg/dL to 0.72 mg/dL in Amblema representing a

67.4 % increase. In Quadrula spp., phosphorus values increased from 0.45 mg/dL at baseline to 0.61mg/dL in November representing a smaller increase of 35.3% compared to A. plicata. Phosphorus levels in A. plicata continued to increase in February and were significantly higher than Quadrula spp. median values at that time (p=0.031). By June, phosphorus levels appeared to return to those similar to baseline for both genera.

Glucose Values and trends are presented by genus for consistency (Figure 3.3). Initially, glucose levels increased by 40.2% after translocation into captivity from a baseline value of 2.46 mg/dL to 3.45 mg/dL in late August. By the next sampling point in November, mean glucose levels returned to values similar to baseline in November and February

then dropped by 30% from 2.5 mg/dL to 1.75mg/dL between February and June.

Glucose median levels did not vary significantly by genus at any time point (Table 3.5).

Isoenzymes No consistent trends were observed in values of ALP within Quadrula spp. over time. At each time point, means of ALP were 2 fold higher in Quadrula spp. versus A. plicata on average. Compared to baseline values, ALT in A. plicata dropped by 35% for the first 5 months in captivity then dropped by an average of 56% between November and June. ALT in Quadrula spp. dropped by an average of 20% for the first 5 months in captivity then dropped by an average of 37% between November and June compared to baseline values. Means of AST in A. plicata dropped on average by 60% between baseline and November. AST levels in Quadrula spp. dropped ~1 (U/L) from baseline at

2.66 (U/L) for the first month in captivity, then in February increased to 3.25(U/L), which is approximately 22% higher than the mean at baseline (Table 3.6).

Concentrations for ALP were statistically higher in Quadrula spp. than in A. plicata at baseline, late August, November, and February (p <0.05) (Figure 3.4). Median values of ALT were significantly higher in A. plicata at baseline, early August, and late

August compared to median values in Quadrula spp. (p<0.05). A. plicata median values of AST were significantly higher than Quadrula spp. at baseline (p<0.05) (Table 3.6).

General Health General health exams for the control group were normal throughout the study.

During winter sampling (November and February) captive control animals that were buried within the substrate, usually Quadrula spp., were presumed to be normal and not disturbed for more extensive examination. All captive controls were alive at the end of the study. Final weight and shell length parameters were not statistically different from intake measurements for either the sample group or the captive controls (p>0.05).

Losses One A. plicata died in transport to captivity, leaving 29 animals available for captive sampling. Overall, 14 of the 29 animals in the captive sampling group brought into captivity were lost due to mortalities that occurred in the facility over the year of study, and 7 animals were lost due to adductor muscle tears at sampling. Over 75% of the losses occurred after November, and more than half of the total losses occurred between

February and June. Adductor muscle tears were more prevalent in A. plicata than in

Quadrula spp. There was no significant association between adductor muscle pathology and death. Small needle tracks in the adductor muscle was observed in 10 animals, but at subsequent samplings 7 of the 10 animals had no observable pathology, indicating that the needle tracks appeared to have healed. Of the original 29 animals in the captive sampling group, 15 were alive at the end of the study, representing a survival rate of

51.7% versus 100% survival in the control group.

3-4 Discussion Previously, only one study has described biochemical parameters in native freshwater mussel hemolymph, and only one species, Elliptio complanata, was included.

This is the first study to evaluate biochemical parameters in the hemolymph of freshwater mussels in captivity over time and the first study to begin to develop reference ranges for

Amblema plicata, Quadrula quadrula, and Quadrula pustulosa. Two genera were included in this study due to a limited number of individuals available in the wild from a single genus or species. In this study, unexpected differences in biochemical parameters between genera at baseline and over time in captivity were found. The results indicate that genus-specific differences in biochemistry of freshwater mussel hemolymph exist.

Further reference ranges may need to be developed for each genus and potentially each species.

Over time both genera show a consistent trend in all electrolyte values (sodium, chloride, magnesium, and potassium) that peaked in November. The sharp increase in values in November, at 5 months post translocation to captivity, could be due to delayed adjustment to captivity, lower filtration rates, decreased ion transport, or other metabolic factors associated with winter quiescence. Circulation of hemolymph is influenced by the cycle of adductor muscle activity, which is also strongly influenced by filtration and feeding activity and seasonal quiescence (Morton 1970). In late November, temperatures are starting to decrease and winter quiescence would have just begun. The spike in electrolytes found in hemolymph in this study may therefore represent the transition period from higher filtration and feeding activities in autumn to lower filtration and feeding activities going into winter that may also influence electrolyte balances in 54

hemolymph. Mean values of phosphorus then peaked in both genera in February at twice the value found in the previous June or subsequent July. Sodium and chloride appear to decrease in the first 14-30 days post translocation, while magnesium and potassium levels were relatively unchanged.

Increased phosphorus levels in water are used as an indicator of water clarity and silt load (Ellison and Brett 2006). Dissolved phosphorus levels in freshwater streams are primarily influenced by fertilizer run off and sewage output (Ellison and Brett 2006).

The Scioto River is an urban water source that also services agricultural lands upriver from the captive facility used in this study. Run-off primarily occurs in the Scioto River between April and May due to snow melt and spring rain; however, levels of phosphorus in hemolymph are similar to baseline levels in June, potentially indicating that the facility filters dissolved nitrates and phosphates effectively or that the values in hemolymph, which could have been higher during the April-May period, have decreased by June.

Particulate phosphorus level increase during siltation events, and conditions such as slow water movement, lower temperatures, and high and low pH values exacerbate particulate phosphorus release from silt (Jitts 1958). Filtration screens small enough to reduce fine particulate matter from entering the facility would also limit food size availability for mussels and such small screens are easily occluded. The facility may be effective at reducing dissolved solutes, but the level of particulate filtration may be ineffective at reducing fine particulate accumulation in animal tanks. A. plicata sits higher in the substrate naturally than Quadrula spp. (normally buried in the substrate), especially in winter. A. plicata is therefore more exposed to fine silt that was observed to

lie over the top of mussel beds during times of low flow in the winter. Interestingly, A. plicata had significantly higher hemolymph phosphorus values than Quadrula spp. in

February. It is unknown if increased phosphorus levels in hemolymph could be detrimental, but it is well documented that silt has detrimental effects on wild freshwater mussel populations (Bilotta and Brazier 2008) primarily due to decreased filtration rate with increased particulate size (Hornbach et al. 1984). Further research is required to determine if high phosphorus levels in hemolymph could be detrimental and if phosphorus levels in hemolymph may be an effective way to quantify silt exposure.

Calcium in bivalves is involved in shell formation of adults and glochidia, acid- base regulation, and respiratory health (Pekkarinen 1997). Calcium moves through the shell to the extrapallial fluid then to hemolymph where Ca+ can enter from the external environment through the gills, intestines, or mantle cavity epithelium. Overall, the system is poised for equilibrium and movement of calcium between compartments (shell, extrapallial fluid, hemolymph, and tissue) is continuous (Comibra et al.1993). The results of this study also showed that calcium is very tightly regulated in this population with little fluctuation over time until the June data. Previous research has documented an increase in calcium levels in hemolymph in response to hypoxic conditions in which calcium is mobilized from the extrapallial fluid to be exchanged for hydrogen ions to maintain ionic equilibrium and hemolymph pH (Comibra et al. 1993). Peak levels of calcium in the hemolymph and extrapallial fluid are also documented in the summer when water temperatures, growth, and metabolism are the highest (Pekkarinen 1997).

Nemczok and Szasz (1975) have also suggested that hemolymph calcium concentrations

in mussels may depend on levels of calcium in water that forms their environment. The last data point in this study was taken in June during expected ―peak times‖ in which the mussels should be undergoing growth and shell formation. Consequently, a decrease in calcium levels in the population implies a general trend of displacement of calcium beyond the reserves of the individual animals potentially due to sustained anoxic conditions, shell formation, or a lack of calcium available in the water in the captive flow through system at that time. Calcium is easily tested and supplemented in aquariums and the results of this study may reflect a need to monitor and potentially supplement calcium levels in captivity during times of peak growth and reproductive activity.

Glucose levels in hemolymph of bivalves are expected to fluctuate seasonally and generally in response to any environmental challenge requiring more energy (Gustafson et al. 2005b). Glucose levels in this study were relatively constant over time except for a small spike in glucose occurring in late August one month post translocation which corresponded with seasonal rises in glucose documented in other species (Pekkarinen

1997) but could also have been in response to acclimation to captivity. These results potentially indicate that handling stress and the stress of translocation into captivity is limited or minimal in this population.

This is the first study to document measurable levels of alanine aminotransferase

(ALT) and alkaline phosphatase (ALP) in freshwater mussels. Gustafson et al.2005, found measurable levels of aspartate aminotransferase (AST) in freshwater mussels that were correlated with mussel size. Alkaline phosphatase (ALP) is an isoenzyme that is usually indicative of liver disease but elevations can occur in response to corticosteroid

activity or exposure to anticonvulsant medications in mammalian species. However,

ALP is also found in seminal fluid of a multitude of mammalian species and is thought to play a role in sperm glycolytic activity and production of fructose (Mann 1964 in Turner and McDonnell 2003). Alanine aminotransferase (ALT) is nonspecific and can be related to liver cell damage or muscle damage (Latimer et al.1994). ALT may be useful in monitoring the effect of repeated sampling on the adductor muscle, however, further study and development of normal versus abnormal levels of isoenzymes is needed to determine their significance in freshwater mussels.

The cause of the losses that occurred during this study is not definitively known.

Up to 5 months in captivity in November, the survival rate was 90%. A quarter of the losses in the sampling group occurred after the November sampling point when animals would normally be burrowed and undisturbed, which is in contrast to the control group that had a 100% survival rate and were neither sampled nor removed from the substrate overwinter. The largest majority of losses and mortalities occurred between February and June, during a time when the metabolic demands increase with warming water temperatures. During November and February, water temperatures in the animal tanks were consistently lower than 12° Celsius, which has not been factored into previous studies in either the wild or captive animals. Cope’s study on the effects of in situ relocation in 2003 showed that doubling or tripling the density of mussels at existing mussel beds did not adversely affect resident or relocated populations. The maximum density that a captive facility can support has not been evaluated in the literature, but nutritive stress has been postulated to contribute to animal losses in extended periods in

captivity (Patterson et al. 1997). Overall, repeated sampling, handling, and the energy required to re-burrow may have decreased nutritive stores and resiliency going into the spring thaw. Further studies are needed to determine if low sampling temperatures, combined with handling or nutritive stress could predispose animals to a higher risk of morbidity or mortality.

3-5 Conclusion This study provides reference ranges for three species of freshwater mussel,

Amblema plicata, Quadrula quadrula and Quadrula pustulosa that will aid in interpretation of future values from the same species. Significant differences between genera, A plicata and Quadrula spp. A greater percentage of losses were seen after winter sampling potentially indicating that animals should not be handled or sampled over winter quiescence in captivity, at least until further studies can determine if overwinter losses are due to the length of time in captivity or winter quiescence. There also appears to be increases in electrolyte values in all animals included in the study in

November, which could also be a result of decreased filtration rates transitioning into winter quiescence. Sodium and chloride decreased and glucose increased in hemolymph in the first month of captivity. These parameters may be indicative of acute stress, requiring additional study. Overall, this study provided a foundation for development of broader reference ranges for these species and a greater understanding of potential challenges in translocation and management of populations in captivity.

Chapter 3 Tables and Figures

Parameter Min Median Mean Max SE Stdev Q1 and Q3 Chloride (mmol/L) 10.5 18.5 17.3 21.4 0.79 3.65 13.5 20.5 Phosphorus (mg/dL) 0.3 0.4 0.4 0.7 0.01 0.09 0.4 0.5 Sodium (mmol/L) 11.0 19.0 18.2 23.0 0.78 3.60 14.5 21.5 Potassium (mmol/L) 0.2 0.5 0.5 0.9 0.04 0.18 0.38 0.6 Magnesium (mg/dL) 0.9 1.3 1.3 2.2 0.08 0.37 1.0 1.5 Calcium (mg/dL) 14.4 19.7 20.8 27.9 0.78 3.58 18.3 22.1 Glucose (mg/dL) 1.0 2.0 2.7 7.0 0.39 1.82 1.0 4.0 ALT (U/L) 0.0 12.0 12.3 33.0 1.52 6.95 10.0 13.5 AST (U/L) 0.0 3.0 5.5 30.0 1.39 6.35 3.0 5.5 ALP (U/L) 0.0 1.2 1.8 9.0 0.68 3.11 1.2 3.9 Weight(g) 231.8 334.8 339.1 410.0 12.6 48.8 320.4 381.7 Length (mm) 93.5 113.3 112.3 126.6 2.05 7.9 107.63 117.7

Table 3.2: Hemocyte biochemical parameters and initial capture measurements from Quadrula quadrula, n=13 and Quadrula pustulosa, n=5. The first quartile shows that 25% of the data are less than or equal to this value. The third quartile shows that 75% of the data are less than or equal to this value. Parameter Min Median Mean Max SE Stdev Q1 and Q3 Chloride (mmol/L) 10.8 14.0 17.33 16.2 0.79 1.55 12.6 14.9 Phosphorus (mg/dL) 0.3 0.45 0.45 0.6 0.02 0.09 0.4 0.5 Sodium (mmol/L) 11.0 15.1 14.7 18.0 0.57 2.40 12.8 16.3 Potassium (mmol/L) 0.1 0.3 0.3 0.4 0.02 0.07 0.26 0.4 Magnesium (mg/dL) 1.1 1.7 1.7 2.5 0.09 0.39 1.4 2.0 Calcium (mg/dL) 14.6 19.6 19.4 23.6 0.68 2.89 16.8 22.2 Glucose (mg/dL) 1.0 2.0 2.2 4.0 0.22 0.92 1.8 3.0 ALT (U/L) 2.0 6.5 7.2 18.0 0.99 4.19 4.0 9.5 AST (U/L) 0.0 2.5 2.7 6.0 0.29 1.23 2.0 3.0 ALP (U/L) 0.0 2.5 3.0 11.0 0.75 3.18 1.0 4.0 Weight(g) 87.9 184.4 176.5 272.7 16.3 60.9 115.0 224.2 Length (mm) 59.8 83.8 81.1 105.5 3.82 14.28 66.3 91.8 60

Electrolyte levels Potassium Magnesium Sodium (mmol/L) Chloride (mmol/L) (mmol/L) (mg/dL) Quadrula Quadrula Quadrula Quadrula A. plicata A. plicata A. plicata A. plicata Genus spp. spp. spp. spp. Mean 14.7 18.2 13.7 17.3 0.3 0.5 1.7 1.3 July 2008 Median 15.5 19.0 14.0 18.5 0.3 0.5 1.7 1.3 Stdev 2.4 3.6 1.6 3.7 0.1 0.2 0.4 0.4 Mean 11.1 10.5 9.9 9.2 0.3 0.3 1.9 1.4 August, Median 11.5 10.0 10.1 9.0 0.2 0.2 2.0 1.3 2wks Stdev 2.2 1.1 1.6 0.9 0.2 0.1 0.4 0.4 Mean 11.1 11.6 11.2 11.2 0.3 0.4 1.5 1.9 August, Median 11.0 11.0 10.0 10.3 0.3 0.4 1.5 1.5 4wks Stdev 1.3 1.6 2.4 2.5 0.1 0.1 0.3 1.2 Mean 17.6 14.8 14.7 13.5 0.5 0.7 2.8 2.2 November Median 19.0 14.0 14.7 13.6 0.5 0.6 2.9 2.1 Stdev 3.3 2.9 0.9 0.8 0.1 0.2 0.5 0.5 Mean 11.6 13.9 12.5 12.7 0.3 0.4 1.7 1.7 February Median 10.0 15.0 12.1 12.6 0.3 0.4 1.7 1.6 Stdev 4.4 3.0 1.7 0.6 0.1 0.2 0.4 0.5 Mean 13.1 9.6 13.3 10.7 0.2 0.2 2.3 1.7 June 2009 Median 15.0 10.0 13.8 11.0 0.3 0.2 2.2 1.6 Stdev 3.1 0.8 2.3 0.9 0.1 0.1 0.6 0.7

Table 3.4: Calcium and phosphorus levels in Quadrula spp. and Amblema plicata at each time point. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9. Calcium and Phosphorus levels Calcium (mg/dL) Phosphorus (mg/dL) Quadrula Quadrula Genus A. plicata A. plicata spp. spp. Mean 19.4 20.4 0.45 0.43 July 2008 Median 19.5 19.7 0.45 0.40 Stdev 2.9 3.6 0.90 0.08 Mean 20.6 20.0 0.48 0.42 August, Median 20.4 21.3 0.50 0.40 2wks Stdev 2.7 2.5 0.14 0.08 Mean 21.3 19.7 0.48 0.58 August, Median 20.4 19.6 0.45 0.50 4wks Stdev 2.3 1.9 0.22 0.18 Mean 20.9 20.7 0.61 0.72 November Median 21.2 21.4 0.60 0.60 Stdev 3.43 2.2 0.10 0.18 Mean 20.2 20.5 0.63 0.78 February Median 20.2 20.2 0.60 0.70 Stdev 2.2 2.2 0.14 0.15 Mean 18.2 18.2 0.55 0.47 June 2009 Median 18.5 17.2 0.50 0.40 Stdev 1.6 2.5 0.16 0.09

Table 3.5: Glucose in Quadrula spp. and Amblema plicata at each time point. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9. Glucose levels Glucose (mg/dL) Quadrula Genus A. plicata spp. Mean 2.2 2.7 July 2008 Median 2.0 2.0 Stdev 0.9 1.8 Mean 2.9 3.1 August, Median 3.0 3.0 2wks Stdev 0.9 1.1 Mean 3.6 3.3 August, Median 3.0 3.0 4wks Stdev 1.5 0.7 Mean 2.8 3.0 November Median 3.0 3.0 Stdev 1.2 0.9 Mean 2.6 2.6 February Median 2.0 3.0 Stdev 1.5 1.0 Mean 1.7 1.9 June 2009 Median 2.0 2.0 Stdev 0.7 0.7

Table 3.6: Isoenzyme levels in Quadrula spp. and Amblema plicata at each time point. Number of each genus sampled at each time point: July A. plicata=21 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4 wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=11, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=9. Isoenzyme levels ALP (U/L) ALT (U/L) AST (U/L) Quadrula Quadrula Quadrula Genus A. plicata A. plicata A. plicata spp. spp. spp. Mean 3.0 1.8 7.2 12.3 2.7 5.5 July 2008 Median 2.5 0.0 6.5 12.0 2.5 3.0 Stdev 3.2 3.1 4.2 7.0 1.2 6.4 Mean 3.2 1.9 6.2 7.9 1.7 1.9 August, Median 2.0 0.0 4.5 7.0 1.0 2.0 2wks Stdev 5.1 4.8 5.7 3.9 2.0 0.8 Mean 2.4 0.9 4.6 8.1 1.6 1.9 August, Median 1.5 0.0 3.0 7.0 1.0 2.0 4wks Stdev 2.4 2.1 3.6 2.8 1.3 1.3 Mean 3.9 1.2 6.1 7.9 2.3 2.6 November Median 4.0 0.0 6.0 8.0 2.0 3.0 Stdev 3.7 2.9 1.9 2.9 1.2 1.0 Mean 2.1 1.3 4.3 4.7 3.3 1.4 February Median 0.5 1.0 4.0 4.0 1.5 1.0 Stdev 3.3 1.4 2.0 2.8 6.9 1.0 Mean 3.2 0.7 4.7 6.1 1.8 1.7 June 2009 Median 3.0 1.0 4.0 6.0 1.0 2.0 Stdev 3.0 0.8 3.7 2.0 1.4 0.5

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Coimbra, A.M., K.G. Ferreira, P. Fernandes, H.G. Ferreira. 1993. Calcium exchanges in Anodonta cygnea: barriers and driving gradients. J. Comp. Physiol. B. 163: 196-202.

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Hornbach, D.J. C. M. Way, T.E. Wissingl and A.J. Burky. 1984. Effects of particle concentration and season on the filtration rates of the freshwater clam, Sphaerium striatinum Lamarck (Bivalvia: Pisidiidae) Hydrobiologia. 108: 83-96.

Fagerland, M.W. and L. Sandvik. 2009. The Wilcoxon Mann Whitney Test under scrutiny. Statistics in Medicine. 28(10): 1487-1497.

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4 Chapter 4: Hematological assessment of freshwater mussels, A. plicata and Quadrula spp.

Abstract Health monitoring that is non-invasive, repeatable, and sensitive is a critical need for North America’s most endangered animal, the freshwater mussel. Ohio is home to a large proportion of these threatened animals and in response has established a captive flow through facility for research and propagation. Currently there is gap in knowledge of mussel physiology and health requirements in the wild and in captivity, representing an urgent need for health assessment methods. In mammalian medicine, a complete blood cell count is a crucial piece of the minimal health database for diagnosis of potential disease or routine health checkups. Recent development of a standardized method for hemocyte characterization in freshwater mussels has paved the way for application of these techniques to freshwater mussels in the wild and in captivity. This study aimed to provide preliminary baseline reference ranges for hemocytology from two common freshwater mussel genera, Amblema and Quadrula in the wild during the peak summer field season and then to translocate animals into captivity to monitor hemocytology trends over one year. Cell differentials were found to be genus-specific at baseline and for the first month in captivity (p< 0.05). Total hemocyte counts between genera differed at 2wks and in November (p <0.10). Four cell types were identified in each genus studied at each time point. Eosinophilic granulocytes predominated in both genera. In A. plicata, eosinophilic granulocytes ranged from 53 to73%, large

agranulocytes from 19 to41%, basophilic granulocytes from 1 to 6% and small agranulocytes from 1 to 3%, compared to Quadrula spp. that had 44-61% eosinophilic granulocytes, 8-27% basophilic granulocytes, 28-40% large agranulocytes, and less than

1 % small agranulocytes. This study provides a foundation for baseline reference ranges for A. plicata and Quadrula spp. and a preliminary understanding of shifts in blood cells in a population of mussels in captivity over one year.

4-1 Introduction Freshwater mussels are centrally located within the aquatic ecosystem, making them highly sensitive and valuable to the ecosystem’s functional health. Freshwater mussel populations have declined due to habitat destruction, pollution, and invasive species invasion and are now critically endangered. Health monitoring that is non- invasive, repeatable, and sensitive is needed. In mammalian medicine, a complete blood cell count is a crucial piece of the minimal health database for diagnosis of potential disease or routine health checkups. In some cases, changes in mammalian blood cell counts are the first indication of disease (Latimer et al.1997). Marine bivalve research has determined that molluscan blood cells, called hemocytes, may respond in a fashion similar to vertebrate blood cells (Hughes et al. 1990), and that changes in hemocyte composition may be indicative of shifts in physiological condition (Ford 1986; Feng

1965). Recent studies on freshwater mussels have shown that hemolymph can be safely and repeatedly drawn, optimally processed, and the cellular portion can be systematically identified and enumerated (Burkhard et al. 2009, Gustafson et al. 2005a). Cellular

reference ranges specific to freshwater mussels are needed to interpret and identify normal and abnormal changes in cell types. The ultimate goal is that cellular data can be used to safely sample a population and gain crucial information about its health without impacting population numbers.

Bivalve immunity is composed of hemocytes and effector molecules that interact to maintain active surveillance through its cellular and innate humoral components.

Circulating hemocytes play a key role in the internal cellular defense mechanism that recognizes and removes non-self particles and in the general maintenance of the internal processes of the animal (Pipe et al. 1997). Phagocytic action is the known primary mechanism of hemocyte defense, with a measurable increase in the number of circulating hemocytes preceding phagocytosis, specifically granulocytes (Pipe and Coles 1995).

Hemocytes are also involved in many internal processes related to physiological vitality, such as gametogenesis, nutrient excretion, digestion, and shell growth and repair (Ford et al. 1993). Bivalves do not have a complement system or memory cells; therefore, observations are truly indicative of real-time challenges. Consequently, monitoring shifts in the quantity and type of certain cells can serve as a sensitive indicator of physiological status, vitality, and immune function.

Hemocyte morphological descriptions for bivalves have been limited to a general classification of granulocyte or agranulocyte. Improved visualization of individual hemocytes began with the incorporation of transmission and scanning electron microscopy starting in the late 1970s and mid 1980s (Rassmussen et al. 1985). However, even with electron microscopy, cell aggregation limited development of a true

morphological schematic and enumeration of a representative circulating population of cells. In addition, methods based on electron microscopy had limited utility for biologists and veterinarians because such equipment is often expensive and impractical for use in the field.

Efforts to make a simple blood smear were also complicated by differential adherence of cell types. Granulocytes adhere and spread on glass, whereas agranulocytes show limited adherence abilities and confirmed that counts performed from hemolymph smears were much lower than counts from cytospins (Lopez et al. 1997). Recently, with the discovery of a novel anticoagulant and the use of cytocentrifugation, a working schema for cellular classification was developed in freshwater mussels (Burkhard et al.

2009). Cytocentrifugation uses a low-speed centrifugal force to separate and deposit a monolayer of cells on slides while maintaining cellular integrity, which ensures that such a monolayer includes granulocytes and agranulocytes. L-cysteine added to cells just prior to cytocentrifugation ensures that cells are adequately separated for more accurate identification.

Bivalves are highly responsive to fluctuating environmental conditions such as pollutants (Pipe et al. 1995), temperature (Feng 1965), and seasonal cycles such as reproduction which affect total cell counts in hemolymph (Suresh and Mohandas 1990).

The logistics of a full year of data collection in the wild is challenging for investigators, due to varying water levels and temperatures, limiting data collection to summer seasons only. Laboratory study can be done year round, but the results are limited in their application back to the population of interest. Use of a captive flow-through system that

attenuates temperature and dynamic fluctuations in ammonia and other run off pollutants may be the best way to begin to develop reliable reference ranges for freshwater mussels that are more inclusive of its year round physiology.

It is our goal to develop a greater understanding of freshwater bivalve hemocytology and a more inclusive reference range for hemocyte differentials by sampling a cohort of freshwater mussels in the wild to establish baseline levels and then translocating them into a captive flow through system for sampling over one year.

Individuals were allowed to acclimate for 14 days then hemolymph samples were drawn in captivity at 2 weeks, 1 month, 5 months, 8 months and 11 months. Total hemocyte counts and cell differentials were performed at each time point. To monitor potential changes in general health post translocation into captivity health measures were observed at each sampling in captivity.

4-2 Materials and Methods

Study Design Forty wild mussels from two genera, Quadrula and Amblema, and three species—

Q. quadrula, Q. pustulosa, and A. plicata—were captured on July 23 2008 at Muskingum

River, Devola, OH. Forty of these subjects were successfully sampled (an adequate amount of hemolymph was obtained) and included in the baseline data group.

Thirty subjects of the original forty were transported into captivity for repeated sampling. Additionally, nine animals were transported into captivity for observation but were not sampled or handled for the duration of the study. The captive control group

consisted of 5 A. plicata, 3 Q. quadrula, and 1 Q. pustulosa. The captive sampling group was comprised of 16 A. plicata, 10 Q. quadrula, and 4 Q. pustulosa. Animals were allowed to acclimate for 14 days prior to sampling. Animals were sampled in early

August, late August, November, February, and June. At each time point the other nine animals were observed for a general health exam only. Mussel health was evaluated by maintenance of position within the substrate, visualization of the ability to retract and/or extend the foot, adductor muscle response to mantle cavity touch, and resistance to manual opening of the shell. Any animal that was unable to exhibit the characteristics above for more than 12-24 hours were presumed dead and removed from the collection and placed into fixative (10% buffered formalin, Fisher Scientific, USA). Animals that had tears in the adductor muscle were not sampled at that time point due to concerns in exposure of the sinus to fluid other than hemolymph and to minimize further damage.

Specimen Collection and Processing Approximately 900µl of hemolymph were collected from each mussel.

Hemolymph was drawn with a 1ml syringe with a 25 G 5/8‖ needle from a sinus in the anterior adductor muscle, as previously described (Gustafson et al. 2005 & Burkhard et al. 2009). Immediately after collection in the field, each sample was transferred to a 1ml microcentrifuge tube at ambient temperature. Hemolymph samples were transported at ambient temperature and processed within 45-60 minutes after collection. Immediately prior to cell counts and slide preparation, samples were individually vortexed for 30 seconds to mechanically separate cells and increase the number of cells in suspension.

Cell counts were prepared as follows: a small aliquot of 20µl of hemolymph from each sample was added at a 1:1 ratio with trypan blue exclusion stain (Mp Biomedicals Inc.,

Solon, OH, USA). The number of live versus dead cells (trypan blue negative and positive, respectively) was enumerated using a Neubauer haemocytometer under light microscopy using 10µl of sample. The total cell count was calculated based on the dilution factor. For slide preparation: an aliquot of 50µl of hemolymph was placed in a cytocentrifuge cup with 50µl of L-cysteine (Calbiochem, EMD Biosciences Inc., La

Jolla, CA, USA) and 10µl of 10% fetal bovine serum (FBS) in phosphate buffered saline

(FBS). Cells were then cytocentrifuged at 0.1 x g for 5 minutes (Shandon Cytospin 4

Cytocentrifuge, Thermo Electron Corporation, Reading, UK). Slides were allowed to air- dry and then stained routinely with Wright’s-Geimsa stain by hand in the field, or by an automated slide-stainer (Harleco Midas2 Automatic Stainer, EMD Chemicals, USA) in the laboratory.

Husbandry All animals were housed at the Columbus Zoo and Aquarium’s Freshwater

Mussel Conservation and Research Center (FMCRC). Water quality parameters and mortalities were recorded weekly. All mussels were kept in 4 x 3 x 2ft tanks with 8-12 inches of sand and gravel for substrate. The system contains a constant total water volume of ~10,000 gallons with a water surface area of 495 sq ft. The number of animals housed in the facility ranges from 1000-2000, depending on the time of year. The facility is a flow through system that utilizes water from the Scioto River pumped through a large

6 ft tower with biological filtration. The facility has the ability to change to a re- circulating system if needed. The average pH of the water during this study ranged from

7.5-8.3 with an osmolality of 25mmol/kg. The pH and hardness varies with natural fluctuations in the river, and is not modified by the system. Ammonia in the Scioto River ranges from 0.001 to 0.8 ppm depending on rain events or run-off from agricultural lands up river. During environmental events that could compromise the water quality (high ammonia concentration or known pollutants) the river water is re-circulated within the initial collection tower up to 3 times to effectively reduce ammonia to tolerable levels between 0.0 and 0.05ppm before contact with the animal collection. The average water and ambient temperature in the facility throughout the year was 17ºC with lows of 6 ºC recorded over winter. Dissolved oxygen was consistently above 95% saturation throughout the study.

Statistical Analysis General descriptive trends for differential cell counts at baseline and over one year are presented. (Evaluation of temporal variability using advanced statistical methods was complicated because the data were dependent (paired) and nonparametric.)

Genus differences appeared to exist in differential cell counts in the baseline data, which was an unexpected result. To determine if these differences were statistically significant, the Mann-Whitney test was used to evaluate differences in differential cell count medians between Quadrula spp. and Amblema plicata at baseline and each subsequent time point, a p-value <0.05 was considered significant (Fagerland and Sandvik 2009). Quadrula

pustulosa and Quadrula quadrula were pooled for comparisons against Amblema due to the low numbers of Quadrula pustulosa (n<5) included in the study. General descriptive trends for cellular differentials over one year are presented. An association between adductor muscle pathology (including both tears and needle tracks) and death as an outcome was evaluated using a Chi square measure of association test with a Yates correction. Initial and final shell length and weight of the control group and the sample group were analyzed using paired t-tests. Statistical analysis was performed using

Minitab 16 Statistical Software (2010) (State College, PA: Minitab, Inc. USA).

4-3 Results

Reference Ranges Reference ranges for cellular differentials in freshwater mussel hemolymph are presented for each genus, due to statistical differences between genera that were found unexpectedly at baseline for eosinophilic granulocytes and basophilic granulocytes

(p<0.05). Consequently, reference ranges for hemocyte differentials in freshwater mussel hemolymph are presented for each genus. (Tables 4.1 and 4.2)

Hemocyte Differential Four cell types were identified in each genus studied at each time point (Figure

4.1 and Table 4.3). ―Early captivity‖ refers to values taken from the first month in captivity from July to Late August. ―Late captivity‖ is defined as the period from

November to June. Eosinophilic granulocytes were the most predominant cell type in 79

each species at baseline and in captivity, ranging from 53 to 73% in A. plicata and 47-

61% in Quadrula spp. A. plicata had a significantly higher proportion of eosinophilic granulocytes at baseline and in late August than Quadrula spp. (p=0.010, p=0.008). The second most predominant cell type in A. plicata was the large agranulocyte, ranging from

19 to 41% over time, followed by a small proportion of basophilic granulocytes (~1-6%) and small agranulocytes (~ 1-3%). A. plicata had a significantly higher proportion of large agranulocytes at 2 weeks post translocation than Quadrula spp. (p=0.027). The second most predominant cell type varied in Quadrula spp., between basophilic granulocytes and large agranulocytes at different time points. For the first month in captivity, the proportion of basophilic granulocytes was 2 – 3 times higher (~21-27%) than the proportion found in the wild in July at baseline (8.6%) in November (9.8%) and in February(8.4%). Quadrula spp. had a significantly higher median percentage of basophilic granulocytes than A. plicata at baseline, early captivity (p=0.000, p=0.000, p=0.007) and at the final time point in June (p=0.086). Large agranulocytes in Quadrula spp. are the second most predominant cell type at baseline (35.5%) and in late captivity

(~28-40%). The percentage of large agranulocytes is significantly higher in Quadrula spp. than in A. plicata at 2 weeks post translocation (p=0.027). In early captivity, a smaller proportion of large granulocytes are represented in Quadrula spp., with population averages of 17-19% at 2 weeks and 4 weeks. Percentages of small agranulocytes in both species are less than 1% in early captivity and in November, but both species show a 2-3 fold increase in the percentage of small agranulocytes in

February at 8 months post translocation. No differences in the median percentage of

small agranulocytes were observed between genera at any time point. Cell differentials were found to be genus-specific at baseline and for the first month in captivity, early and late August respectively (p< 0.05) (Figure 4.1).

Total Hemocyte Count Total hemocyte counts between genera differed significantly in early August and in February (p <0.10) (Figure 4.2 and Table 4.4). Quadrula spp. had consistently higher total hemocyte counts than A. plicata at each time point. From baseline to November,

Quadrula spp. had, on average, 33-38 % more hemocytes than A. plicata. Total hemocyte counts were significantly higher in Quadrula spp. than in A. plicata in early

August 2 weeks post translocation and in February, 8 months post translocation (p=0.057, p=0.098).

General Health General health exams for the control group were normal throughout the study.

During winter samplings (November and February) captive control animals that were buried within the substrate were presumed to be normal and not disturbed for more extensive examination. All captive controls were alive at the end of the study. Final weight and shell length parameters were not statistically different from intake measurements for either the sample group or the captive controls.

February and June. Adductor muscle tears were more prevalent in A. plicata than in

51.7% versus 100% survival in the control group.

4-4 Discussion

Cellular Differentials This study showed that a recently described cellular schema for freshwater mussels (Burkhard et al. 2009) can be used successfully to describe four distinct cell types not only for Quadrula spp. but also for Amblema plicata. This study is the first study to develop baseline reference ranges for Amblema plicata and Quadrula quadrula, and Quadrula pustulosa.

Two genera were included in this study due to a limited number of individuals available in the wild from a single genus or species. In this study, unexpected differences in cellular differential counts between genera at baseline and over time in captivity were found. The results indicate that genus specific differences in cell types in freshwater mussels exist. Further cellular reference ranges may need to be developed for each genus and potentially each species.

To date the link between cellular morphology and cellular function is not yet defined in freshwater mussels. Previous work in bivalve hemocytology has been limited to describing differing percentages of granulocytes and agranulocytes involved in phagocytic, chemokinetic, or cytotoxic activity (Friebel and Renwrant 1995). Marine bivalve research in commercially important clams, mussels, and oysters have reported general trends in cell function among granulocytes and agranulocytes. In marine bivalves, granulocytes appear to be more common than agranulocytes, making up more than half of cell samples regardless of species (Cima et al. 2000, Foley and Cheng 1975,

Pipe et al. 1997, Cheng 1983). Both granulocytes and agranulocytes are capable of phagocytosis and secretion of humoral factors, but only a small percentage of agranulocytes of the total population have been observed engaging in phagocytosis. The majority of actively phagocytic granulocytes have eosinophilic granules (Cima et al.

2000, Tripp 1992, Foley and Cheng 1975). This study also observed eosinophilic granulocyte differential prevalence of over 50% in both species throughout the study.

Basophilic granulocytes were present in higher proportions in Quadrula spp. during the first month in captivity potentially representing a cell type that may function separately

from eosinophilic granules potentially similar to neutrophils and macrophages in mammalian cells. Agranulocytes have been found in oysters in greater number in circulation during times of intense tissue damage and inflammation post infection with invasive parasites (Ford et al. 1993, Santarem et al. 1994). Interestingly, the present study found a higher proportion of small and large agranulocytes between February and

June during a time when the total number of animals with adductor mussel pathology was at its highest. While this study cannot directly link cell types with cell functions and causative agents, it provides a strong preliminary foundation of reference ranges and trends for future studies. The ability to identify specific cell types will hopefully lead to expanded knowledge of cellular function in the future.

Total Hemocyte Counts Hemocytes are considered the foundation of the bivalve immune defense system; therefore, quantification of hemocytes has historically been used as a general measure of health in marine species. The results of this study reveal that a subtle increase in total hemocyte count (roughly 20%) was observed in both genera between 1 month and 5 months in captivity and a precipitous decline in hemocyte counts were observed in

February in both genera. The causes of the observed increases and subsequent declines in hemocyte concentration in the sample population are not definitively known. In previous work, increases in total hemocyte counts have most commonly been attributed to contaminant exposure and have served as a general indicator of cellular stimulation and migration in experimental laboratory settings (Pipe and Coles 1995; Suresh &

Mohandas 1990). Such increases in total cell count may result from proliferation of cells or from movement of the cells from the tissues into circulation. Decreases in hemocyte counts in bivalves have been documented in response to starvation (Butt et al. 2007), shell repair (Allam et al. 2000), and reproduction (Sarterem et al. 1994; Suresh and

Mohandas 1990). Decreases could also be a consequence of lysis, decreased recruitment, or from movement of cells from circulation into the tissues.

Fluctuations in hemocyte numbers can be due to numerous internal and external causes that could be indicative of normal physiological changes or pathological states.

For example, Suresh and Mohandas (1990) reported that hemocytes in the gonads mobilize to clean up cellular debris in the post spawning period and are thus absent in circulation, resulting in low hemocyte counts during the regression period. Similar results were reported by Sanarem et al. (1994), finding that their lowest hemocyte counts occurred in July during the post-spawning period and a time of low food availability due to their utility in internal processes. However, natural external factors also influence hemocyte counts Feng (1965) reported that hemocyte counts in oysters had a positive linear relationship with heart rate and ambient temperature. At ambient temperatures of

6º, 12º, 18º, 22ºC, hemocyte counts were found to be 1.6, 2.7, 4.1 and 4.9 million respectively. The corresponding heart rates were 5, 9, 19 and 26 beats per minute.

Bayne (1973) also reported results confirming the effect of ambient temperature on hemocyte enumeration. Circulation of cells is also influenced by the cycle of adductor muscle activity, which is, in turn, also strongly influenced by filtration and feeding activity and seasonal quiescence (Morton 1970). Consequently, depressed cell counts in

February may simply be explained by ambient temperature drops and relative quiescence due to temperatures below 10ºC on average in the facility during that month.

This study found a large amount of variation in hemocyte counts, with standard deviations of greater than 90% of the mean and high standard errors of the mean at each time point. Hemocyte adherence to collection tubes and formation of large aggregates of cells can falsely decrease cell counts. Based on findings by Burkhard (2009) cells were enumerated after partial re-suspension by vortexing in an attempt to mechanically separate cells. Vortexing prior to loading hemolymph in the hemocytometer appeared to partially re-suspend cells, resulting in small clumps that could still be enumerated.

Fixation has been used most frequently to minimize aggregation. However, intense degranulation in response to fixatives have been shown to obscure visualization of hemocytes due to an increase in cellular debris as well as falsely skew enumeration of granulocytes in the total count (Cheng and Foley 1974, Sanarem et al. 1994).

Losses Animals collected for this study were from apparently healthy individuals that maintained general measures of health throughout translocation and transition into captivity for the first 5 months. Losses occurred primarily during winter: times when animals would normally be burrowed and undisturbed. Repeated sampling combined with overall handling and the energy required to re-burrow may have decreased resiliency. Without a true wild population control group or a control group kept within a

tighter temperature regime, it is unknown if captivity or temperature effects contributed to the declines.

4-5 Conclusions This research provides a foundation of cellular reference ranges for Amblema plicata and Quadrula spp. Cell differentials were found to be genus-specific at baseline and for the first month in captivity. Total hemocyte counts between genera differed significantly in early August and in February. Further investigation into potential variation in freshwater hemolymph cellular composition by season, species, and housing is needed. Further physiological studies linking cellular function with cellular morphologies and expansion of broader reference ranges, perhaps by season, are next steps toward utility of this diagnostic method for interpretation. Losses that occurred during the winter period may indicate a need to refrain from sampling during times of low temperature.

Chapter 4 Tables and Figures

Table 4.1: Hemocyte differential count and total cell count/ml from 22 Amblema plicata mussels from the Muskingum River, Devola, Ohio captured in June. Quartile 1 represents the first quartile in which 25% of the data are less than or equal to this value. Q3 represent the third quartile in which 75% of the data are less than or equal to this value. Hemocyte Type Min Median Mean Max SE Stdev Q1 Q3 Eosinophilic 21.0 79.0 73.2 92.0 3.6 16.84 63.3 84.3 Granulocyte Basophilic 0.0 0.0 0.7 6.0 0.3 1.45 0.0 1.0 Granulocyte Large 8.0 19.5 25.0 73.0 3.4 15.96 14.8 34.8 Agranulocyte Small 0.0 0.0 1.0 6.0 0.3 1.51 0.0 2.0 Agranulocyte Total Cell Count 6,000 55,000 69,000 164,000 10,095 47,352 30,500 114,500

Table 4.2: Hemocyte differential count and total cell count/ml from 13 Quadrula quadrula and 5 Quadrula pustulosa mussels from the Muskingum River, Devola, Ohio captured in June. Quartile 1 represents the first quartile in which 25% of the data are less than or equal to this value. Q3 represent the third quartile in which 75% of the data are less than or equal to this value. Hemocyte Type Min Median Mean Max SE Stdev Q1 Q3 Eosinophilic 0.0 60.0 54.9 87.0 6.04 25.63 44.0 74.8 Granulocyte Basophilic 0.0 6.0 8.6 21.0 1.72 7.32 3.0 18.3 Granulocyte Large 5.0 30.5 35.6 100.0 6.32 26.80 30.5 46.8 Agranulocyte Small 0.0 1.000 0.89 3.000 0.227 0.963 0.000 2.000 Agranulocyte Total Cell Count 12,000 65,000 94,000 354,000 21,625 91,746 36,500 118,500

Granulocyte % Granulocyte % Agranulocyte % Agranulocyte % Quadrula Quadrula Quadrula Quadrula A. A. plicata A. plicata A. plicata Genus spp. spp. spp. spp. plicata Mean 54.9 68.7 8.6 0.7 35.6 29.8 0.9 0.7 July 2008 Median 60.0 78.0 6.0 0.0 30.5 22.0 1.0 0.0 Stdev 25.6 19.7 7.3 1.8 26.8 18.3 1.0 1.1 Mean 59.1 67.5 21.2 1.1 19.1 31.6 0.6 0.4 August, Median 54.0 71.0 14.5 1.0 16.5 29.0 0.0 0.0 2wks Stdev 14.5 16.9 14.4 1.6 9.7 17.1 1.3 0.5 Mean 54.7 71.7 27.1 2.6 17.8 25.1 0.4 0.5 August, Median 62.0 73.0 24.5 1.0 18.0 20.0 0.0 0.0 4wks Stdev 18.4 13.6 21.3 6.3 9.3 15.5 0.9 0.7 Mean 61.9 61.5 9.9 5.9 27.6 32.5 0.6 0.1 November Median 58.0 60.0 6.0 1.0 25.0 33.0 0.0 0.0 Stdev 14.0 14.0 10.3 12.2 14.5 8.8 1.2 0.3 Mean 45.0 53.7 8.0 3.1 43.9 40.6 3.1 2.6 February Median 45.0 56.0 4.0 2.0 45.0 38.0 2.0 1.0 Stdev 17.1 15.6 9.2 3.8 14.2 14.0 3.9 4.3 Mean 47.8 54.9 15.2 6.4 36.0 36.9 1.0 1.9 June 2009 Median 38.0 67.0 10.0 3.0 37.0 32.0 1.0 1.0 Stdev 19.9 26.7 16.1 12.7 25.7 22.8 1.3 2.0

Table 4.4: Total cell counts (cells/ml) represented in Quadrula spp. and Amblema plicata at each time point. Number of each genus sampled at each time point: July A. plicata=22 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=14, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=10. Total cell count Total cells/mL Quadrula Genus A. plicata spp. Mean 94,000 53,273 July 2008 Median 65,000 36,000 Stdev 91,746 41,302 Mean 98,571 55,067 August, Median 73,000 42,000 2wks Stdev 72,411 26,075 Mean 89,286 67,067 August, Median 89,000 68,000 4wks Stdev 44,539 23,975 Mean 108,246 82,831 November Median 85,600 76,000 Stdev 86,222 58,238 Mean 60,500 14,857 February Median 34,000 12,000 Stdev 64,544 5,398 Mean 109,556 41,429 June 2009 Median 96,000 42,000 Stdev 80,745 14,820

Figure 4.1: Eosinophilic granulocytes, basophilic granulocytes, large agranulocytes, and small agranulocytes in Quadrula spp. and Amblema plicata. Baseline Total cell counts per ml of hemolymph for each genus is listed for reference: Quadrula spp.:94,000 cell/ml mean, 65,000 cells/ml median. Amblema plicata: 53,273 cells/ml mean, 36,000 cells/ml median. Asterisks represent statistically significant differences between genera at that time point (p<0.05) except basophils in June (p-value <0.10). Medians reported by a solid dot. Means and standard error brackets reported. Black bars marked with hatch marks represent baseline values taken from the animals in the wild. Note: the scale for small agranulocytes is 10%. Number of each genus sampled at each time point: July A. plicata=22 Quadrula spp.=18, Aug 2wks A. plicata=15 Quadrula spp.=14, Aug 4wks A. plicata=15 Quadrula spp.=14, November A. plicata=13 Quadrula spp.=14, February A. plicata=7 Quadrula spp.=12, June A. plicata=7 Quadrula spp.=10.

References

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Bayne, B.L. 1973. Physiological changes in Mytilus edulis L. induced by temperature and nutritive stress. Journal of Marine Biological Association of the United Kingdom. 53:39- 58.

Butt, D., S. Aladaileh, W.A. O'Connor and D.A. Raftos. 2007. Effect of starvation on biological factors related to immunological defence in the Sydney rock oyster (Saccostrea glomerata). Aquaculture. 264:82-91.

Cheng, T.C. 1988. In vivo effects of heavy metals on cellular defense mechanisms of Crassostrea virginica:Total and differential cell counts. Journal of Invertebrate Pathology. 51:207-214.

Cheng, T.C. and D.A. Foley. 1975. Hemolymph cells of the bivalve mollusk Mercenaria mercenaria: An electron microscopic study. Journal of Invertebrate Pathology. 26:341- 351.

Cima, F, V. Matozzo, M. G. Marin, and L. Ballarin.2000. Hemocytes of the clam Tapes philippinarium(Adams & Reeve, 1850): morphofunctional characterization. Fish and Shellfish Immunology. 10:677-693.

Fagerland, M.W. and L. Sandvik. 2009. The Wilcoxon Mann Whitney Test under scrutiny. Statistics in Medicine. 28(10): 1487-1497.

Feng, S.Y. 1965. Heart rate and leukocyte circulation in Crassostrea virginica. Biological Bulletin. 128:198-210.

Foley D.A, and T.C Cheng. 1974. Morphology, hematologic parameters, and behavior of hemolymph cells of the Quahog clam, Mercenaria mercenaria. Biological Bulletin. 146:343-356.

Foley D.A, and T.C Cheng. 1975. A quantitative study of phagocytosis by hemolymph cells of the pelecypods Crassostrea virginica and Mercenaria mercenaria. Journal of Invertebrate Pathology. 25:189-197. 93

Ford, S.E. 1986. Effect of repeated hemolymph sampling on growth, mortality, hemolymph protein and parasitism of oysters, Crassostrea virginica. Comparative Biochemical Physiology Part A. 85(3):465-470.

Ford, S.E., S.A Kanaley, and D.T.J. Littlewood. 1993. Cellular responses of oysters infected with Haplosporidium nelsoni: changes in circulating and tissue-infiltrating hemocytes. Journal of Invertebrate Pathology. 61:49-57.

Friebel, B. and L. Renwrantz. 1995. Application of density gradient centrifugation for separation of eosinophilic and basophilic hemocytes from Mytilus edulis and characterization of both cell groups. Comparative Biochemical Physiology Part A. 112(1):81-90.

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Lopez, C., M.J. Caballal, C. Azevedo, and A. Villalba. 1997. Morphological characterization of the hemocytes of the clam, Ruditapes decussates (Mollusca: Bivalvia). Journal of Invertebrate Pathology. 69:51-57.

Pipe, R.K., Farley, S.R. and J.A. Coles. 1997. The separation and characterization of haemocytes from the mussel Mytilus edulis. Cell Tissue Research. 289:537–545.

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5 Chapter 5: Conclusion

This thesis aimed to 1) elucidate the optimal methods to process and transport hemolymph in order to improve cell enumeration and minimize in-vitro effects and maintain cellular viability for further physiological assessment; and 2) apply developed diagnostic techniques to establish baseline reference ranges for hematologic and chemical parameters of hemolymph and 3) follow trends over time in captivity. Conclusions have been generated which contribute to the field of freshwater mussel conservation and aquatic medicine.

5-1 Practical guidelines on how to transport and process hemolymph in order to preserve cellular integrity . Hemocytes in vitro were found to be sensitive to changes in pH and should be

processed at a pH level most similar to physiological levels in their home

environment.

. Freshwater mussel hemocytes should be transported untreated at ambient

temperature and ideally processed within 4 hours.

. L-cysteine (25mg/ml) reconstituted at an optimal pH of for the animal should

be added to hemolymph just prior to preparation of slides for cellular

differentials.

5-2 Chemical and hematological baseline reference ranges for freshwater mussels . Chemical and hematological parameters varied significantly by genus between

Amblema plicata and Quadrula spp.

. Broader sampling is needed to ensure their application outside of the home

watershed and season collected, but results provide a preliminary foundation

that can be applied forward to mussels of the same species.

5-3 Trends in chemical and cytological parameters in freshwater mussels housed in a captive flow through facility over one year . A greater percentage of losses were seen after sampling during winter versus

captive controls that were not handled or sampled, potentially indicating that

animals should not be handled or sampled winter quiescence.

. Declines in sodium, chloride, and potassium were seen in both genera in the

first month in captivity.

. All electrolyte values in both genera had marked increases in November

potentially due a low filtration period and the beginning of winter quiescence.

. Calcium and glucose values remained steady in the population until the last

collection point in June when metabolic demands were higher with warmer

temperatures and values in both parameters declined, potentially indicating a

time when more hands on management is needed to support populations.

The techniques applied and guidelines developed can be adapted to additional freshwater species in captivity and in the wild. Such methods will be useful to other researchers studying endangered or threatened freshwater mussel species. As more researchers utilize these techniques valuable reference ranges can be built upon to assist in interpretation for a larger variety of species. Furthermore, as invertebrates are the taxon showing the highest sensitivity to environmental disturbance, with the development of more sensitive measures of health we will be better able to evaluate our restoration efforts in streams and lakes where they reside, benefiting the aquatic ecosystem and the terrestrial species, including us, that rely on it.

References

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