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EXTRA-RENAL MECHANISMS OF OSMOTIC AND ACID/BASE REGULATION IN A EURYHALINE ELASMOBRANCH (Dasyatis sabina)

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EXTRA-RENAL MECHANISMS OF OSMOTIC AND ACID/BASE REGULATION IN A EURYHALINE ELASMOBRANCH (Dasyatis sabina)
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2008

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Chlorides ( jstor )
Epithelium ( jstor )
Excretion ( jstor )
Fish ( jstor )
Fresh water ( jstor )
Gills ( jstor )
Ions ( jstor )
Plasmas ( jstor )
Salinity ( jstor )
Sea water ( jstor )
St. Johns River, FL ( local )

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University of Florida
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University of Florida
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Copyright the author. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
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5/4/2003
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70786466 ( OCLC )

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EXTRA-RENAL MECHANISMS OF OSMOTIC AND ACID/BASE REGULATION IN A EURYHALINE ELASMOBRANCH ( Dasyatis sabina ) By PETER MARC PIERMARINI A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2002

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This dissertation is dedicated to the memories of Dr. Thomas B. Thorson, for his seminal research on the osmoregulatory physiology of bull sharks in Lake Nicaragua; Dr. Homer W. Smith, for his enthusiastic spirit of adventure in comparative physiology; and William Bartram, whose appreciation and love of Florida’s wildlife has inspired generations.

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ACKNOWLEDGMENTS I would like to first thank the chair of my doctoral committee, Dr. David H. Evans for advising me during this project and providing me with the opportunity to pursue my interests in elasmobranch physiology. Dave has helped me develop into a confident, independent scientist for which I am extremely grateful. Dave has also unselfishly prioritized my needs and interests over his own many times, which speaks volumes of his dedication to his graduate students. My accomplishments would not have been possible without his guidance and generosity. I also am grateful for the assistance and guidance provided by the other current and former members of my doctoral committee: Drs. Lauren Chapman; Louis Guillette, Jr.; Karl Karnaky, Jr.; Frank Nordlie; and Jill Verlander. Lauren Chapman’s expertise in ecology and statistics helped me analyze and interpret my results in ways that are more unique and usually more appropriate than ways found in the general comparative physiology literature. I greatly appreciate the generosity of Lou Guillette for allowing me to use equipment and supplies in his laboratory. Also, interactions with Lou and his graduate students (especially Ed Orlando and Andy Rooney) were extremely helpful and integral to my progress as a graduate student. Karl Karnaky’s enthusiasm for and positive attitude toward science are examples for every scientist to follow. I would like to thank Karl for his interest in my project and for his mentorship at the Mount Desert Island Biological Laboratory during the summer of 1998. I’d like to thank Frank Nordlie for allowing me to use equipment in his lab. Frank is one of the most genuine people that iii

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I met during my graduate career and I greatly enjoyed my interactions with him. I am extremely thankful to Jill Verlander for serving as the external member of my committee. For an “external” member, Jill has had the most influence on the direction of my dissertation and I tried to model my approach to understanding epithelial function after her own. I greatly appreciate all of the time she volunteered to teach me new techniques and to honestly discuss my results. I would also like to express my appreciation of support and help provided by other colleagues, faculty, graduate students, and undergraduates, especially Elena Amesbury, Craig Aubrey, Alex Bond, Sarah Bouchard, George Burgess, Keith Choe, Noah DeVincente, Kirk Giesbrandt, Sarjeet Gill, Amy Gilliam, Linda Green, Mark Gunderson, William Harvey, Mike Janech, Harvey Lillywhite, Steve McCormick, Mike Miyamoto, Ed Orlando, Suhel Quader, Nicole Reid, Rob Robins, Diana Rosman, Ines Royaux, Buck Snelson, Laura Sirot, Colette St. Mary, Marianne Thompson, Leigh Truong, and Steve Walsh. I also received excellent logistical support from the staff of the Zoology Fiscal Office, secretaries of the Zoology Main Office, staff of the Zoology Stockroom, and staff of the Seahorse Key Marine Laboratory. My research was primarily supported by an EPA STAR Fellowship (U-915419-01-0), with partial funding provided by Sigma Xi, Theodore Roosevelt Memorial Fund, International Women’s Fishing Association, American Elasmobranch Society, and the Explorer’s Club. Partial funding of my research was also provided by David Evans (NSF Grants IBN-9604824, IBN-9306997, and IBN-0089943) and by the Department of Zoology. My research project would not have been possible without animal holding iv

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facilities and vehicles provided by the Department of Zoology, as well as facilities provided by the University of Florida Seahorse Key Marine Laboratory. I thank all my close family and friends for their support throughout my graduate education, especially my parents and grandparents. Without their understanding, encouragement, and financial assistance, this would have been a much longer and difficult road for me to take. Lastly I would like to thank Laura Sirot for her love and companionship these past 6 years, which have made my life a lot more enjoyable. v

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TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iii LIST OF TABLES.............................................................................................................ix LIST OF FIGURES.............................................................................................................x ABSTRACT......................................................................................................................xii CHAPTERS 1 GENERAL INTRODUCTION......................................................................................1 Introduction...................................................................................................................1 Extra-Renal Ion Regulatory Mechanisms in Marine Teleosts......................................2 NaCl Transport......................................................................................................2 Acid/Base Transport..............................................................................................6 Extra-Renal Ion Regulatory Mechanisms in Marine Elasmobranchs...........................8 NaCl Transport......................................................................................................8 Acid/Base Transport............................................................................................12 Extra-Renal Ion Regulatory Mechanisms in Freshwater Teleosts..............................13 NaCl and Acid/Base Transport............................................................................13 Extra-Renal Ion Regulatory Mechanisms in Freshwater Elasmobranchs...................17 NaCl and Acid/Base Transport............................................................................17 Overview of Dissertation Research............................................................................19 2 OSMOREGULATION OF ATLANTIC STINGRAYS FROM THE FRESHWATER ST. JOHNS RIVER, FLORIDA..................................................................................25 Introduction.................................................................................................................25 Materials and Methods................................................................................................27 Field Studies........................................................................................................27 Acclimation Experiments....................................................................................28 Statistical Analyses..............................................................................................30 Results.........................................................................................................................31 Plasma Osmolytes and Rectal Glands of Freshwater Atlantic Stingrays............31 Acclimation of Freshwater Atlantic Stingrays to Increased Salinity..................31 Discussion...................................................................................................................32 vi

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Osmoregulation of Freshwater Atlantic Stingrays..............................................32 Acclimation of Freshwater Atlantic Stingrays to Higher Salinities....................35 Evolutionary and Ecological Implications..........................................................36 3 EFFECT OF ENVIRONMENTAL SALINITY ON NA+,K+-ATPASE IN THE GILLS AND RECTAL GLAND OF THE ATLANTIC STINGRAY........................42 Introduction.................................................................................................................42 Materials and Methods................................................................................................46 Animal Collection and Holding Conditions........................................................46 Collection of Tissues...........................................................................................47 Na+,K+-ATPase Activity.....................................................................................48 Immunoblots........................................................................................................49 Immunohistochemistry........................................................................................51 Statistical Analyses..............................................................................................53 Results.........................................................................................................................53 Na+,K+-ATPase Activity.....................................................................................53 Immunoblots........................................................................................................54 Immunohistochemistry........................................................................................54 Discussion...................................................................................................................55 Gills.....................................................................................................................55 Rectal Gland........................................................................................................58 4 IMMUNOCHEMICAL ANALYSIS OF THE VACUOLAR PROTON-ATPASE B-SUBUNIT IN THE GILLS OF THE ATLANTIC STINGRAY: INFLUENCE OF SALINITY AND RELATION TO NA+,K+-ATPASE................................................66 Introduction.................................................................................................................66 Materials and Methods................................................................................................69 Animal Collection and Holding Conditions........................................................69 Collection of Gill Tissue.....................................................................................70 Antibodies...........................................................................................................71 Immunoblotting of V-H-ATPase B-subunit........................................................71 Immunohistochemical Localization of V-H-ATPase B-subunit.........................74 Colocalization of V-H-ATPase with Na+,K+-ATPase.........................................75 Statistical Analyses..............................................................................................75 Results.........................................................................................................................76 Immunoblotting of V-H-ATPase B-subunit........................................................76 Immunohistochemistry of V-H-ATPase B-subunit.............................................76 Colocalization of V-H-ATPase Immunoreactivity with Na+,K+-ATPase...........77 Discussion...................................................................................................................77 5 IMMUNOCHEMICAL EVIDENCE OF A PENDRIN-LIKE CL-/HCO3EXCHANGER IN THE GILL EPITHELIUM OF THE ATLANTIC STINGRAY...90 Introduction.................................................................................................................90 Materials and Methods................................................................................................92 vii

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Animal Collection and Holding Conditions........................................................92 Collection of Gill Tissue.....................................................................................93 Antibodies...........................................................................................................94 Immunoblot Analysis of Pendrin Immunoreactivity...........................................95 Immunohistochemical Localization of Pendrin Immunoreactivity.....................97 Colocalization of Pendrin Immunoreactivity with V-H-ATPase and Na+,K+-ATPase................................................................................................................98 Statistical Analyses..............................................................................................99 Results.........................................................................................................................99 Immunoblot Analysis of Pendrin Immunoreactivity...........................................99 Immunohistochemical Localization of Pendrin Immunoreactivity.....................99 Colocalization of Pendrin Immunoreactivity with V-H-ATPase and Na+,K+-ATPase..............................................................................................................100 Discussion.................................................................................................................100 6 SUMMARY AND FUTURE DIRECTIONS............................................................114 Summary...................................................................................................................114 Future Directions......................................................................................................117 What is the Function of the Rectal Gland in Freshwater Atlantic Stingrays?...118 What is the Mechanism of Branchial Na+/H+ Exchange?.................................119 How Can the Proposed Model of Branchial Ion Regulation Be Tested?..........119 What Controls Expression of Ion Regulatory Mechanisms?............................120 Is the Proposed Model Applicable to Other Elasmobranchs?...........................121 What Limits Elasmobranch Euryhalinity?........................................................122 LIST OF REFERENCES.................................................................................................124 BIOGRAPHICAL SKETCH...........................................................................................146 viii

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LIST OF TABLES Table page 2-1 Comparison of osmolality and osmolytes in freshwater Atlantic stingray plasma to Lake Jesup water, other freshwater elasmobranchs, a freshwater teleost, and marine Atlantic stingrays........................................................................................................38 2-2 Comparison of osmolality and osmolytes in ambient water and plasma osmolality of Experimental Atlantic stingrays.................................................................................39 2-3 Comparison of plasma osmolality and osmolytes in Control group (Day 32) and Field group.................................................................................................................40 ix

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LIST OF FIGURES Figure page 1-1 Current model of active NaCl secretion in the marine teleost chloride cell and marine elasmobranch rectal gland tubule cell........................................................................22 1-2 Current model of acid/base excretion in the seawater teleost gill..............................23 1-3 Current model of active NaCl uptake and acid/base excretion in the freshwater teleost gill...................................................................................................................24 2-1 Osmotic composition of plasma from Control and Experimental groups on Days 12, 22, and 32...................................................................................................................41 3-1 Activity of Na+,K+-ATPase in the gills and rectal gland of freshwater, seawater-acclimated, and seawater Atlantic stingray................................................................61 3-2 Representative immunoblot for Na+,K+-ATPase in gill and rectal gland membrane enrichments of freshwater, seawater-acclimated, and seawater stingrays.................62 3-3 Relative abundance of immunoreactivity for 111 or 112 kDa band representing the -subunit of Na+,K+-ATPase in gills and rectal gland of freshwater, seawater-acclimated, and seawater stingrays............................................................................63 3-4 Representative photomicrographs of Na+,K+-ATPase immunostaining in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays........................................................................................................64 3-5 Number of Na+,K+-ATPase-rich cells in the gills of freshwater, seawater-acclimated, and seawater stingrays................................................................................................65 4-1 Representative immunoblot for V-H-ATPase B-subunit in gill membrane protein from freshwater, seawater-acclimated, and seawater Atlantic stingrays....................83 4-2 Relative abundance of immunoreactivity for 60.5 kDa protein representing the B-subunit of V-H-ATPase in the gill membrane enrichments of freshwater, seawater-acclimated, and seawater stingrays............................................................................84 4-3 Representative photomicrographs of V-H-ATPase immunostaining in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays........................................................................................................85 x

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4-4 Number of V-H-ATPase-rich cells in the gills of freshwater, seawater-acclimated, and seawater Atlantic stingrays..................................................................................86 4-5 Representative higher magnification photomicrographs of V-H-ATPase-rich cells in gills from freshwater, seawater-acclimated, and seawater Atlantic stingrays............87 4-6 Representative photomicrographs of V-H-ATPase immunolabeling colocalized with Na+,K+-ATPase in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays...............................................................88 4-7 Hypothetical model of NaCl and acid/base transport in the gills of the Atlantic stingray.......................................................................................................................89 5-1 Representative immunoblot for pendrin immunoreactivity in gill membrane protein from freshwater, seawater-acclimated, and seawater Atlantic stingrays..................105 5-2 Relative abundance of immunoreactivity for 144 kDa band representing pendrin, in gill membrane protein of freshwater, seawater-acclimated, and seawater Atlantic stingrays....................................................................................................................106 5-3 Representative photomicrographs of pendrin immunolabeling in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays......................................................................................................107 5-4 Higher magnification photomicrographs of pendrin immunolabeling in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays......................................................................................................108 5-5 Representative photomicrographs of pendrin immunolabeling colocalized with V-H-ATPase in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays.............................................................109 5-6 Higher magnification photomicrographs of pendrin immunolabeling colocalized with V-H-ATPase in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays.............................................................110 5-7 Representative photomicrographs of colocalization of pendrin immunolabeling colocalized with Na+,K+-ATPase in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays..........................111 5-8 Higher magnification photomicrographs of pendrin immunolabeling colocalized with Na+,K+-ATPase in longitudinal sections of gill filaments from freshwater, seawater-acclimated, and seawater Atlantic stingrays.............................................................112 5-9 Model of NaCl and acid/base regulation in the freshwater and seawater elasmobranch gill.....................................................................................................113 xi

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy EXTRA-RENAL MECHANISMS OF OSMOTIC AND ACID/BASE REGULATION IN A EURYHALINE ELASMOBRANCH (Dasyatis sabina) By Peter M. Piermarini May 2002 Chair: Dr. David H. Evans Department: Zoology The Atlantic stingray is unique because it is one of the few elasmobranchs to establish populations in both fresh and seawater environments. Before this dissertation, the mechanisms that allow elasmobranchs to live in such contrasting environments were unknown. The goals of this dissertation were to describe the general osmoregulation of the Atlantic stingray, and to establish the extra-renal mechanisms this species uses for osmotic and acid/base balance in fresh and seawater environments. First, I described the osmoregulatory strategy of freshwater Atlantic stingrays. I established that the plasma of freshwater stingrays had relatively low urea and NaCl concentrations, and a small salt-secreting rectal gland, compared to marine Atlantic stingrays. When freshwater stingrays were acclimated to seawater, plasma urea and NaCl concentrations increased to typical seawater Atlantic stingray levels, which suggested that the stingrays were not physiologically restricted to freshwater environments. xii

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Second, I investigated the influence of salinity on Na+,K+-ATPase expression in gills and rectal glands of Atlantic stingrays. In gills, Na+,K+-ATPase expression (activity, immunoreactivity, and number of Na+,K+-ATPase-rich cells) was highest in freshwater stingrays, compared to seawater individuals. In rectal glands, Na+,K+-ATPase activity and immunoreactivity were higher in seawater stingrays, compared to freshwater individuals. These results suggested that the gills are important for active ion uptake in fresh water, while the rectal gland is important for active ion secretion in seawater. Third, I focused on expression of vacuolar-proton-ATPase (V-H-ATPase) in stingray gills. The V-H-ATPase immunoreactivity was higher in gills from freshwater stingrays, compared to seawater individuals. Localization of V-H-ATPase was basolateral in relatively large cells of the gill epithelium that were not Na+,K+-ATPase-rich. I proposed that V-H-ATPase-rich cells were sites of Cl-/HCO3exchange and that Na+,K+-ATPase-rich cells were sites of Na+/H+ exchange. Last, I described the expression of a pendrin-like transporter in the Atlantic stingray gill. Pendrin is a Cl-/HCO3exchanger that plays an important role in HCO3excretion in the mammalian kidney. Pendrin immunoreactivity was highest in gills of freshwater stingrays and occurred in the apical region of V-H-ATPase-rich cells. This suggested that V-H-ATPase-rich cells are sites of Cluptake and HCO3excretion in stingray gills. xiii

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CHAPTER 1 GENERAL INTRODUCTION Introduction The osmoregulatory physiology of most fishes is specialized for life in a particular environment (e.g., fresh or seawater), but many species can live in a relatively wide range of salinities (Evans 1984b). These euryhaline fishes are of great physiological interest, because they can differentially express their ion regulatory mechanisms depending on the environment they occupy. Among the more important ion regulatory processes in euryhaline fishes are those responsible for NaCl and acid/base homeostasis. Although the kidneys of fishes play an important role in water and divalent ion balance, extra-renal organs such as gills and salt glands are the primary sites of NaCl and acid/base regulation. Many studies have investigated the cellular mechanisms of NaCl and acid/base regulation in the gills of marine and freshwater teleosts (bony fishes; e.g., trout, salmon, killifish, tilapia), which has been aided by the fact that many teleost families contain species that are euryhaline (Evans 1984b). In contrast, data are lacking on the cellular mechanisms associated with extra-renal ion regulation in marine and freshwater elasmobranchs (cartilaginous fishes; e.g., sharks, skates, rays). Elasmobranchs primarily inhabit marine environments, and euryhaline species are uncommon. Of the approximate 1000 elasmobranch species, only 43 are found in freshwater environments (Compagno and Cook 1995). Furthermore, only 14 of these 43 species are considered euryhaline; the remainder are stenohaline freshwater stingrays of the family Potamotrygonidae (Compagno and Cook 1995). 1

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2 The Atlantic stingray (Dasyatis sabina) is an exceptional elasmobranch, because it is one of the few species that can live and reproduce in both marine and freshwater environments. It is a common, coastal species that ranges from the Chesapeake Bay to Central America (Bigelow and Schroeder 1953), and is known to enter freshwater rivers during summer (Gunter 1938, Sage et al. 1972, Schwartz 1995). In the St. Johns River of Florida, the Atlantic stingray has established populations over 300 km from the Atlantic Ocean that reproduce and complete their life cycle in fresh water (Johnson and Snelson 1996), which makes them the only resident freshwater elasmobranch population in North America. Therefore, it is possible to obtain freshwater and marine individuals for comparative studies. In addition, the Atlantic stingray’s small body size and abundance relative to other elasmobranchs make it an ideal species to study elasmobranch extra-renal NaCl and acid/base regulatory mechanisms. The goals of this dissertation are to describe the general osmoregulation of the Atlantic stingray, and establish some of the extra-renal mechanisms it uses to maintain NaCl and acid/base balance in both marine and freshwater environments. In this chapter I summarize the known extra-renal ion regulatory mechanisms of teleost and elasmobranch fishes in marine and freshwater environments, and provide an overview of the research described in the following chapters. Extra-Renal Ion Regulatory Mechanisms in Marine Teleosts NaCl Transport The plasma of marine teleosts is hypoosmotic and hypoionic to seawater (Karnaky 1998), which leads to an osmotic loss of water and a diffusional influx of ions (primarily Na+ and Cl-), respectively. To prevent dehydration, marine teleosts drink seawater (Smith 1930, Potts and Evans 1967) and absorb the ingested NaCl across their intestine,

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3 which allows water to follow osmotically (Loretz 1995). Therefore, marine teleosts require effective NaCl excretory mechanisms to offset the diffusional and ingested NaCl loads. The primary site of net NaCl excretion is the gills (Evans 1979); the teleost kidney cannot produce a urine that is hyperosmotic, hypernatric, or hyperchloric to the plasma (Smith 1930, Hickman and Trump 1967). The gill epithelium is composed of three main cell types that are directly or indirectly involved with ion regulation: 1) pavement cells; 2) mitochondrion-rich chloride cells; and 3) accessory cells. Pavement cells are the predominant cell type in the gills, and compose over 90% of the branchial surface epithelium (Laurent and Perry 1995). Compared to other cells in the gills, pavement cells are squamous in shape, contain few mitochondria, and have numerous apical microridges that are hypothesized to enhance gas exchange (Laurent 1984, Laurent and Perry 1995). Important to ion regulation, pavement cells form deep tight junctions (many interconnected strands) with adjacent cells (pavement, chloride, or accessory), and these junctions are impermeable to ions (Sardet et al. 1979, Sardet 1980, Karnaky 1992). Chloride cells are large, ovoid cells primarily found between lamellae on the trailing (afferent) edge of gill filaments (Laurent and Dunel 1980, Laurent 1984, Karnaky 1986, Pisam and Rambourg 1991, Van Der Heijden et al. 1997). These cells were first identified by Keys and Wilmer (1932) and are the cellular sites of active NaCl excretion in the gill epithelium (Foskett and Scheffey 1982). Chloride cells are characterized by large numbers of mitochondria; a branching, cytoplasmic tubular system formed by extensive basolateral plasma membrane infoldings; and an abundance of subapical vesicles (Laurent and Dunel 1980, Laurent 1984, Karnaky 1986, Pisam and Rambourg

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4 1991). The infolded basolateral membrane provides numerous binding sites for Na+,K+-ATPase (Karnaky et al. 1976, Hootman and Philpott 1979) and places this active transporter in close proximity to mitochondria. Chloride cells are often found in multicellular groups with other chloride cells and/or accessory cells; the closely associated apical membranes of these cells form a shallow, concave pit or crypt that sinks below adjacent pavement cells (Laurent 1984, Karnaky 1986). In addition, chloride cells form shallow tight junctions (few interconnecting strands) with adjacent chloride or accessory cells, and these junctions are permeable (leaky) to ions (Sardet et al. 1979, Hootman and Philpott 1980, Sardet 1980, Karnaky 1992). Accessory cells are mitochondrion-rich, but usually are smaller in size, have less extensive basolateral membrane infoldings, and have a lower Na+,K+-ATPase activity, compared to chloride cells (Hootman and Philpott 1980, Karnaky 1986, Pisam and Rambourg 1991). Accessory cells were hypothesized to be developing or degenerating chloride cells (Sardet et al. 1979, Hootman and Philpott 1980, Wendelaar Bonga and van der Meij 1989, Wendelaar Bonga et al. 1990), but their true identity and function are unknown. Regardless, accessory cells form shallow, leaky tight junctions with adjacent chloride cells, as noted above. Key model tissues that have been crucial to understanding the NaCl excretory mechanisms of the marine teleost gill are the opercular epithelium of Fundulus heteroclitus and Oreochromis mossambicus, and jaw skin of Gillichthys mirabilis. Opercular and jaw skin epithelia are simple, flat, and composed almost entirely of chloride cells that are morphologically identical to those in the gills (Karnaky and Kinter 1977, Marshall and Nishioka 1980, Foskett et al. 1981, Karnaky 1986). Studies on these

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5 epithelial sheets mounted in Ussing chambers, under short-circuit conditions, showed that transport of Clacross the epithelium was active (Degnan et al. 1977, Karnaky et al. 1977, Marshall and Nishioka 1980) and that Clsecretion occurred exclusively in chloride cells (Foskett and Scheffey 1982). No net movement of Na+ was detected under short-circuit conditions (Degnan et al. 1977, Degnan and Zadunaisky 1980, Marshall 1981), which suggested that Na+ secretion was not active and occurred passively through leaky tight junctions between chloride and accessory cells (Degnan and Zadunaisky 1980, Marshall 1981). The model of NaCl secretion by teleost chloride cells was originally proposed by Silva et al. (1977a), and was recently reviewed by Marshall and Bryson (1998) and Evans et al. (1999). According to the model (Figure 1-1), Clsecretion is indirectly energized by a basolateral Na+,K+-ATPase that establishes a favorable gradient for Na+ entry into the cell. This Na+ gradient drives a basolateral Na+K+2Cl-cotransporter that brings 2 Cland 1 K+ into the cell for every Na+. A basolateral Na+,K+-ATPase pumps Na+ back out of the cell, but Claccumulates within the cell and exits passively though an apical Clchannel. This selective secretion of Clcreates an outside-negative electrical potential that pulls Na+ through leaky tight junctions of the paracellular pathway between chloride and accessory cells. Several lines of evidence have verified components of this model. For example, a basolateral Na+,K+-ATPase was shown by experiments where ouabain (a Na+,K+-ATPase inhibitor) significantly decreased branchial and opercular NaCl secretion when applied basolaterally, but not apically (Karnaky et al. 1977, Silva et al. 1977a, Degnan and Zadunaisky 1979). Additionally, and -subunits of Na+,K+-ATPase were cloned from

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6 the gills of a few teleost species (Kisen et al. 1994, Cutler et al. 1995, Hwang et al. 1998), and localization studies have shown that Na+,K+-ATPase activity and immunoreactivity occur on the basolateral membrane of chloride cells (Karnaky et al. 1976, Hootman and Philpott 1979, Witters et al. 1996, Wilson et al. 2000b). Basolateral Na+K+2Cl-cotransporter activity has been identified by studies that inhibited opercular Clsecretion with the loop diuretics furosemide (an inhibitor of NaCl cotransport) and bumetanide (an inhibitor of Na+K+2Clcotransport) (Degnan et al. 1977, Eriksson et al. 1985). Also, recent immunohistochemical studies have localized Na+K+2Cl-cotransporter immunoreactivity to chloride cells (Wilson et al. 2000b, Pelis et al. 2001). An apical Clchannel with functional similarity to the human cystic fibrosis transmembrane regulator (CFTR) was detected by patch-clamp analysis of primary cell cultures from the operculum (Marshall et al. 1995). Additionally, Singer et al. (1998) cloned and sequenced a CFTR homologue from the marine teleost gill that was 59% identical to human CFTR, and a recent immunohistochemical study localized CFTR immunoreactivity to the apical region of chloride cells (Wilson et al. 2000b). Acid/Base Transport Acid/base regulation involves the excretion of excess protons (H+) and/or bicarbonate (HCO3-), which are the respective acidic and basic ions. While both the gills and kidneys contribute to acid/base balance, the gills play the predominant role (Claiborne 1998). However, the mechanisms that marine teleost gills use to maintain acid/base homeostasis have not been studied extensively as those for NaCl excretion. Recent evidence suggests that the chloride cell is the cellular site of acid/base transport mechanisms in the marine teleost gill (see below), but contributions by pavement cells and/or accessory cells cannot be ruled out. The current model of

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7 acid/base transport mechanisms in the gills of marine teleosts is similar to Krogh’s (1939) model of acid/base and NaCl regulation in freshwater fishes, and has been recently reviewed by Claiborne (1998). According to the model (Figure 1-2), apical extrusion of H+ to seawater occurs in exchange for passive, electroneutral Na+ uptake via a Na+/ H+ exchanger (NHE), while HCO3-extrusion occurs in exchange for passive, electroneutral Cluptake via a band-3 Cl-/HCO3anion exchanger (AE-1). A basolateral Na+,K+-ATPase maintains the favorable gradient for Na+ uptake by pumping Na+ into the blood. The fate of the absorbed Clis not known. It is interesting to note that the activity of these apical exchangers would enhance NaCl loading, which is opposite to the animal’s ion regulatory needs in seawater. Therefore, H+ and HCO3secretion may increase the cost of NaCl regulation, although they occur through passive exchangers. Several lines of evidence support some of the components of this model, but not all are unequivocal. The apical NHE has the most experimental support and was identified in studies that showed H+ excretion by marine teleosts was dependent on external Na+ (Evans 1977, Evans 1982, Claiborne et al. 1997) and inhibited by external amiloride (an inhibitor of Na+ exchangers and channels) (Claiborne et al. 1997). Additionally, an apical NHE isoform (NHE-2) was partially sequenced in marine teleost gills (Claiborne et al. 1999), and immunoreactivity for NHE-2 (Wilson et al. 2000b) and NHE-3 (another apical isoform) was localized to chloride cells (Edwards et al. 1999, Wilson et al. 2000b). Apical AE-1 activity was suggested by a study that showed apical HCO3secretion was dependent on external Cland inhibited by external 4,4’-diisothiocyanatostilbene-2,2’-disulfonic acid (DIDS), an inhibitor of anion transporters (Claiborne et al. 1997).

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8 However, no molecular or immunochemical evidence of AE-1 in the marine teleost gill has been published, and the exact cellular location of AE-1 activity is unknown. Extra-Renal Ion Regulatory Mechanisms in Marine Elasmobranchs NaCl Transport The plasma of marine elasmobranchs is slightly hyperosmotic to seawater, because of renal reabsorption of and low branchial permeability to urea and trimethylamine oxide (TMAO); this leads to an osmotic influx of water (Shuttleworth 1988, Karnaky 1998). However, the plasma of marine elasmobranchs is substantially hypoionic to seawater (Shuttleworth 1988, Karnaky 1998), which results in a diffusional influx of NaCl. Therefore, elasmobranchs do not need to drink seawater, but are still required to excrete excess NaCl. The site of net NaCl excretion in marine elasmobranchs is the digestive tract via a specialized salt-secreting rectal gland (Burger and Hess 1960). In contrast to marine teleosts, the gills are considered a site of net NaCl uptake and not NaCl excretion (Bentley et al. 1976), but some evidence challenges this idea (see below). The kidneys of elasmobranchs cannot produce urine that is hyperosmotic, hypernatric, or hyperchloric to the plasma (Burger 1965). The rectal gland has been well established as the primary site of net NaCl excretion in marine elasmobranchs. Burger and Hess (1960) determined that the rectal gland was a key organ for net NaCl excretion, because it secreted a solution that contained twice the NaCl concentration of the plasma. The digitiform rectal gland is found between the spiral valve intestine and rectum, and contains thousands of secretory tubules that empty into a central lumen, which is continuous with the intestine (Hoskins 1917, Shuttleworth 1988, Valentich et al. 1995). The tubule cells are mitochondrion-rich with extensive basolateral membrane infoldings (Bulger 1963) that provide a large surface area for

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9 insertion of Na+,K+-ATPase (Eveloff et al. 1979, Valentich et al. 1995), and also place this transporter in close proximity to mitochondria. Similar to marine teleost chloride cells, tight junctions between adjacent tubule cells are shallow and considered leaky to ions (Ernst et al. 1981, Forrest et al. 1982). Two important experimental approaches that led to the understanding of NaCl secretory mechanisms in the rectal gland were the in vitro perfused gland (Hayslett et al. 1974, Silva et al. 1990) and the isolated, perfused rectal gland tubule (Forrest et al. 1983, Greger and Schlatter 1984b) from the spiny dogfish (Squalus acanthias). Both of these techniques showed that Cltransport across the tubule cells was active and that Na+ transport to the tubule lumen was passive, through leaky tight junctions of the paracellular pathway (Silva et al. 1977b, Silva et al. 1983, Greger and Schlatter 1984b). The current model of NaCl secretion by rectal gland tubule cells was originally proposed by Silva et al. (1977b) and has been reviewed recently by Riordan et al.(1994) and Silva et al.(1997). The model is identical to that of NaCl secretion by the marine teleost chloride cell, except that NaCl is secreted into the tubule lumen instead of the seawater environment (Figure 1-1). Several lines of evidence verified components of this model. A basolateral Na+,K+-ATPase was shown by experiments that inhibited NaCl secretion when ouabain was applied basolaterally to in vitro perfused rectal glands and isolated perfused tubule preparations (Silva et al. 1977b, Greger and Schlatter 1984a). Additionally, cytochemical staining of Na+,K+-ATPase is consistent with a basolateral membrane localization (Goertmiller and Ellis 1976, Eveloff et al. 1979). Basolateral Na+K+2Cl-cotransporter activity was suggested by studies that inhibited Clsecretion of perfused glands and

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10 tubules with furosemide and bumetanide (Silva et al. 1977b, Forrest et al. 1983, Greger and Schlatter 1984b, Greger and Schlatter 1984a, Palfrey et al. 1984), and by experiments that inhibited Cluptake into rectal gland, basolateral membrane vesicles with furosemide and bumetanide (Eveloff et al. 1978, Hannafin et al. 1983, Hannafin and Kinne 1985). A bumetanide-sensitive Na+K+2Clcotransporter has been cloned and sequenced from the rectal gland (Xu et al. 1994) and was immunolocalized to the basolateral membrane of rectal gland tubule cells (Lytle et al. 1992). At least two types of apical Clchannels in rectal gland tubule cells were detected by patch-clamp studies (Greger et al. 1985, Greger et al. 1986, Gogelein et al. 1987, Greger et al. 1987), with one channel having functional similarity to human CFTR (Riordan et al. 1994). A shark homologue of CFTR with 72% amino acid similarity to human CFTR was cloned from the rectal gland and localized to the apical region of rectal gland tubule cells (Marshall et al. 1991, Lehrich et al. 1998). Although the rectal gland is considered the primary site of net NaCl excretion in elasmobranchs, it is important to note that animals with surgically removed or ligated rectal glands still hyporegulate plasma NaCl concentrations (Burger 1965, Chan et al. 1967, Haywood 1975, Evans et al. 1982). This suggests another organ may contribute to NaCl excretion, such as the gills and/or kidneys, but direct physiological or biochemical evidence for such branchial or renal transport has not been published. The elasmobranch gill epithelium is not considered to play a major role in NaCl excretion, but it contains mitochondrion-rich cells in a similar anatomical location as marine teleost chloride cells (Wright 1973, Laurent and Dunel 1980, Crespo 1982, Laurent 1984, Crespo and Sala 1986, Conley and Mallatt 1988, Wilson et al. 1997). Two different mitochondrion-rich cell types were identified in the elasmobranch gill based on

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11 apical membrane morphology (Laurent and Dunel 1980, Crespo 1982, Laurent 1984), but functional or biochemical differences between the two cell types have not been described. Elasmobranch mitochondrion-rich cells are slightly different from marine teleost chloride cells in that basolateral membrane infoldings are much less extensive and do not form a cytoplasmic tubular network (Laurent and Dunel 1980, Laurent 1984, Wilson et al. 1997). Other important differences with marine teleost chloride cells are that elasmobranch mitochondrion-rich cells appear singly in the gill epithelium and always form deep tight junctions with adjacent pavement cells (Karnaky 1998). Accessory cells have never been described in the elasmobranch gill epithelium. Despite their differences, marine teleost chloride cells and elasmobranch mitochondrion-rich cells have some similarities, such as numerous mitochondria, many subapical vesicles, and elevated activity of Na+,K+-ATPase relative to surrounding cells (Wright 1973, Laurent and Dunel 1980, Laurent 1984, Conley and Mallatt 1988). Also, Evans and More (1988) provided evidence of a basolateral bumetanide-sensitive Na+NH4+2Clcotransporter in marine elasmobranch gills, whose function may be similar to that of a Na+K+2Clcotransporter. However, the cellular location of this transporter was not determined, and any further immunochemical and molecular evidence of this transporter in the elasmobranch gill have not been published. Overall, direct physiological, immunochemical, or molecular evidence of NaCl secretory mechanisms in the elasmobranch gill are lacking. A flat epithelium that is analogous to the teleost operculum or jaw skin has not been discovered for the elasmobranch gill, which has limited progress in understanding this tissue’s contribution to NaCl balance.

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12 Acid/Base Transport In contrast to the minor role elasmobranch gills appear to play in active NaCl secretion, they are the predominant site of acid/base related transport in marine elasmobranchs (Heisler 1988). However, the exact cells and cellular mechanisms involved have not been firmly established. Several lines of evidence suggest that acid excretion by the marine elasmobranch gill is mediated by an apical NHE, as in marine teleosts. Results from whole animal flux experiments showed that acid extrusion in elasmobranchs was dependent on external Na+ (Evans et al. 1979, Evans 1982). Additionally, immunoreactivity for apical NHE isoforms (NHE-2 and NHE-3) was detected in the elasmobranch gill (Choe et al. 2002, Edwards et al. 2002) and localized to mitochondrion-rich cells (Edwards et al. 2002). Interestingly, an NHE may not be the only mechanism of H+ secretion, because vacuolar-proton-ATPase (V-H-ATPase) activity and immunoreactivity were also detected in mitochondrion-rich cells of the marine elasmobranch gill (Wilson et al. 1997). In freshwater teleost fishes, V-H-ATPase plays an important role in apical H+ extrusion (see below), and may perform a similar function in the marine elasmobranch gill. The marine elasmobranch gill secretes HCO3(Holder et al. 1955, Swenson and Maren 1987), presumably through an apical Cl-/ HCO3exchanger (Swenson and Maren 1987, Claiborne and Evans 1992). However, sensitivity of HCO3secretion to external Clor DIDS has not been investigated. Also, the exact molecular identity of the HCO3secretion mechanism is not known because no molecular or immunochemical evidence of a Cl-/ HCO3exchanger in the gills of a marine elasmobranch have been published.

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13 Extra-Renal Ion Regulatory Mechanisms in Freshwater Teleosts NaCl and Acid/Base Transport The plasma of freshwater teleosts is hyperosmotic and hyperionic to their environment (Karnaky 1998), which leads to an osmotic uptake of water and a diffusional loss of NaCl, respectively. To prevent volume loading, freshwater teleosts excrete copious amounts of dilute urine (Hickman and Trump 1967, Evans 1979) and substantially lower their drinking rates (Potts and Evans 1967). Diffusional and urinary NaCl losses are offset by active branchial NaCl uptake from fresh water (Perry 1997, Karnaky 1998). In freshwater teleosts, the branchial mechanisms of active NaCl uptake are inextricably linked to those of acid/base regulation. Therefore, these mechanisms are discussed together. The gill epithelium of freshwater teleosts is composed of pavement and chloride cells; accessory cells are not present in the freshwater teleost gill epithelium (Laurent and Dunel 1980, Laurent 1984, Pisam et al. 1990). Most pavement cells of freshwater teleost gills have a similar morphology to those from seawater teleost gills, but several studies have identified a subset of mitochondrion-rich pavement cells that may play an active role in NaCl and acid/base transport (Goss et al. 1992, Goss et al. 1994, Goss et al. 1998, Goss et al. 2001, Galvez et al. 2002). Both types of pavement cells in freshwater teleosts form deep tight junctions with adjacent pavement or chloride cells, and the junctions are impermeable to ions (Sardet et al. 1979, Karnaky 1992). Chloride cells are found between lamellae and also appear sporadically on lamellae in the freshwater teleost gill epithelium. Similar to marine teleosts, chloride cells from freshwater species are mitochondrion-rich, have elaborate basolateral infoldings, and are Na+,K+-ATPase-rich (Laurent and Dunel 1980, Laurent 1984, Pisam and Rambourg

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14 1991, Perry and Laurent 1993, Uchida et al. 1996, Witters et al. 1996, Perry 1997, Wilson et al. 2000a). Ultrastructural studies have suggested that two populations of chloride cells exist in the freshwater teleost gill (Pisam et al. 1987, Pisam et al. 1988, Pisam et al. 1990, Pisam and Rambourg 1991, Pisam et al. 1995), but it is not clear if any biochemical or functional differences exist between these cells. In fact, they may represent different developmental stages of the same cell type (Perry 1997). The apical membrane of freshwater chloride cells is often flush with or extends beyond adjacent pavement cells and forms microridges or microvilli with elaborate shapes and patterns that vary between species (Perry et al. 1992). Chloride cells from freshwater teleost gills always form deep tight junctions with surrounding pavement or chloride cells, and the junctions are impermeable to ions (Sardet et al. 1979, Karnaky 1986, Pisam et al. 1990, Karnaky 1992). The cellular sites of active NaCl uptake and acid/base excretion in freshwater teleost gills are debatable. Chloride cells were suggested to be the sites of NaCl uptake and acid/base excretion based on direct correlations between chloride cell surface area and NaCl uptake (Perry and Laurent 1989, Laurent and Perry 1990). However, more thorough morphological studies provided evidence that Na+/H+ and Cl-/HCO3exchange occurred in pavement and chloride cells, respectively (Goss et al. 1992, Goss et al. 1994, Goss and Perry 1994, Perry and Goss 1994, Goss et al. 1998). The current consensus is that chloride cells are the primary site of Cl-/ HCO3exchange, and that both chloride and pavement cells can contribute to Na+/H+ exchange, depending upon the species (Perry 1997, Wilson et al. 2000a).

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15 One reason for the uncertainty of which cells are involved is that the teleost operculum and jaw skin have not proven to be effective model systems for understanding chloride cell function in freshwater teleosts (Wood and Marshall 1994). The opercular epithelium of freshwater bony fish does not have an abundance of chloride cells (Foskett et al. 1981, Marshall et al. 1997, Burgess et al. 1998), and when mounted in Ussing chambers, the operculum does not actively absorb NaCl (Degnan et al. 1977, Marshall et al. 1997) or the epithelium has extremely low rates of NaCl uptake (Burgess et al. 1998). Another approach to determining the sites of ion uptake in freshwater teleost gills is the use of cell culture to grow flat sheets of gill epithelial cells. Prt et al. (1993) developed a technique to yield primary cultures of gill epithelial cells in freshwater teleosts, but these cultures are plagued by problems concerning the polarity (distinct apical and basolateral membranes) of the cultured cells and the poor survival of chloride cells in culture conditions (Prt and Bergstrom 1995). Recently, chloride cells were successfully added to cultures of pavement cells, but this preparation fails to show active NaCl uptake (Fletcher et al. 2000). The original model of NaCl uptake and acid/base excretion in freshwater teleosts was hypothesized by Krogh (1939) who proposed that Na+ and Cluptake were independent of one another, and were directly linked to electroneutral exchanges of H+ and HCO3-, respectively. Therefore, excretion of a H+ could not occur without simultaneous uptake of Na+. However, Avella and Bornancin (1989) presented data that challenged Krogh’s model and suggested Na+ uptake was energized by electrogenic H+ secretion via a putative H+-translocating ATPase. The current model has been reviewed by Evans et al. (1999) and was modified recently by Wilson et al. (2000a). According to

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16 the model (Figure 1-3), an apical V-H-ATPase on pavement or chloride cells actively excretes H+, which creates a favorable electrochemical gradient for apical Na+ uptake via an epithelial Na+ channel (ENaC). A basolateral Na+,K+-ATPase transports Na+ from the cells to the blood. Uptake of Cland HCO3excretion occur exclusively in chloride cells via an apical, electroneutral AE-1. The mechanism that energizes the apical AE-1 is not known, and a mechanism for basolateral Cltransport from chloride cells to the blood has not been determined. Several lines of evidence support the proposed model. A V-H-ATPase was detected in both pavement and/or chloride cells of freshwater teleosts by immunohistochemistry and in situ hybridization (Lin et al. 1994, Sullivan et al. 1995, Sullivan et al. 1996, Wilson et al. 2000a). Immunocytochemistry showed that V-H-ATPase was localized to the apical plasma membranes of both pavement and chloride cells (Sullivan et al. 1995, Wilson et al. 2000a). Additionally, Fenwick et al. (1999) have shown that Na+ uptake in freshwater teleosts is blocked by external application of bafilomycin (a specific V-H-ATPase inhibitor), and the V-H-ATPase B-subunit has been cloned and sequenced from the gills of a freshwater teleost (Perry et al. 2000). Evidence for an apical ENaC was provided by Clarke and Potts (1998), who showed that external benzamil (a Na+ channel inhibitor) blocked Na+ uptake across the gills, while external application of a specific NHE inhibitor (5-(N,N-dimethyl)-amiloride) had no effect on branchial Na+ uptake. Additionally, Wilson et al. (2000a) localized ENaC immunoreactivity to the apical region of both pavement and chloride cells. In several freshwater teleost species, Na+,K+-ATPase has been immunolocalized to chloride cells (Uchida et al. 1996, Ura et al. 1996, Witters et al. 1996, Schreiber and Specker 1999,

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17 Seidelin and Madsen 1999, Dang et al. 2000b, Wilson et al. 2000a, Zydlewski and McCormick 2001), where it occurs on the basolateral plasma membrane (Dang et al. 2000a, Wilson et al. 2000a). Pavement cells also contain Na+,K+-ATPase activity, but it is much lower than that of chloride cells (Kultz and Jurss 1993, Galvez et al. 2002). Apical Cl-/HCO3exchange activity was detected when an externally applied anion exchanger inhibitor (4-acetamido-4’-isothiocynato-stilbene-2,2’-disulphonic acid) reduced rates of Cluptake (Perry et al. 1981, Perry and Randall 1981). Additionally, Sullivan et al.(1996) showed that chloride cells expressed mRNA for AE-1 using in situ hybridization, and Wilson et al. (2000a) have shown that AE-1 immunoreactivity occurs in the apical region of chloride cells. Extra-Renal Ion Regulatory Mechanisms in Freshwater Elasmobranchs NaCl and Acid/Base Transport The plasma of euryhaline elasmobranchs in freshwater is hyperosmotic and hyperionic to the environment. Although plasma urea and TMAO are reduced to about half that of marine elasmobranchs, the plasma osmolarity of freshwater individuals is almost twice that of freshwater teleosts (Smith 1931, Urist 1962, Thorson 1967, Thorson et al. 1973, Otake 1991), which results in an extremely high osmotic influx of water. To prevent volume loading, freshwater elasmobranchs excrete copious volumes of dilute urine (Smith 1931), with urine flows exceeding those found in freshwater teleosts (Hickman and Trump 1967). In contrast to euryhaline elasmobranchs in fresh water, stenohaline freshwater stingrays of the family Potamotrygonidae no longer retain urea and TMAO in their plasma (Thorson et al. 1967, Thorson 1970, Griffith et al. 1973, Gerst and Thorson 1977, Bittner and Lang 1980), which presumably decreases their osmotic uptake of water.

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18 Regardless of osmoregulatory strategy, freshwater elasmobranchs require mechanisms of NaCl uptake to offset diffusional and urinary losses. It is assumed that the gills are the site of active Na+ and Cluptake, in exchange for H+ and HCO3excretion, respectively; however, no studies on ion regulatory mechanisms in the gills of a freshwater elasmobranch have been published. Pang et al. (1977) described preliminary evidence that suggested Potamotrygonid stingrays extract Na+ from fresh water through an active mechanism, but its sensitivity to inhibitors or exchanges with counter ions (e.g., H+) were not investigated. It is assumed that the gill epithelium of freshwater elasmobranchs is composed of pavement and mitochondrion-rich cells, but histological or ultrastructural studies on the freshwater elasmobranch gill have not been published. Also, the expression of or activity of ion transporters known to occur in gills of marine elasmobranchs (e.g., Na+,K+-ATPase and V-H-ATPase) have not been determined in the gills of freshwater species. Although no research has investigated ion regulatory mechanisms in the gills of freshwater elasmobranchs, a few studies have examined the morphology and biochemistry of rectal glands from freshwater species. As noted above, freshwater elasmobranchs face diffusional losses of NaCl to the environment. Therefore, a salt-secreting rectal gland would appear to have no beneficial ion regulatory function in a freshwater species. As expected, rectal glands from freshwater elasmobranchs are considerably smaller than those from marine species (Thorson et al. 1978, Thorson et al. 1983). Additionally, the reduced size of the gland has been correlated with a smaller number of secretory tubules (Oguri 1964, Thorson et al. 1978) and a lower overall ATPase activity in the tubule cells (Gerzeli et al. 1976).

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19 Overview of Dissertation Research From the above review, it is apparent that the branchial mechanisms of NaCl and acid/base regulation are poorly understood in elasmobranchs, especially in freshwater species. The mechanisms of NaCl secretion by the rectal gland of marine elasmobranchs are well established, but few studies have examined these mechanisms in rectal glands of freshwater species. In the following chapters, I attempt to enhance our understanding of the extra-renal mechanisms (especially branchial) that elasmobranchs use to maintain NaCl and acid/base homeostasis in marine and freshwater environments. In Chapter 2, I describe the osmoregulatory physiology (e.g., plasma osmolytes and rectal gland size) of freshwater Atlantic stingrays from the St. Johns River, and determine if they have the ability to acclimate to seawater. As mentioned previously, this species is considered euryhaline, but no research has determined if the freshwater populations, which reproduce and complete their life cycle in the St. Johns River, have the typical euryhaline elasmobranch osmoregulatory strategy in fresh water. Furthermore, it has not been shown that these freshwater populations still retain the physiological ability to live in seawater. It is possible that during their occupation of the river, they have shown signs of physiological divergence from marine elasmobranchs, as found in freshwater Potamotrygonid stingrays. Results from this chapter were published in the following paper: “Piermarini, P. M. and Evans, D. H. (1998) Osmoregulation of the Atlantic stingray (Dasyatis sabina) from the freshwater Lake Jesup of the St. John River, Florida Physiol Zool 71, 553-560”. In Chapter 3, I measure the activity and expression of Na+,K+-ATPase in the gills and rectal gland of freshwater and marine Atlantic stingrays, and determine what changes in Na+,K+-ATPase activity and expression occur when freshwater stingrays are

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20 acclimated to seawater. This transporter has been studied extensively in teleost gills and plays an important role in NaCl balance of both freshwater and marine teleosts. While Na+,K+-ATPase has been well studied in the rectal glands of marine elasmobranchs, the transporter’s role in freshwater rectal glands or in the gills of marine and freshwater elasmobranchs has not been established. Results from this chapter were published in the following paper: “Piermarini, P. M. and Evans, D. H. (2000) Effects of environmental salinity on Na+/K+-ATPase in the gills and rectal gland of a euryhaline elasmobranch (Dasyatis sabina) J Exp Biol 203, 2957-2966”. In Chapter 4, I compare the expression of V-H-ATPase between the gills of freshwater and marine Atlantic stingrays, determine if V-H-ATPase is expressed by the same cells that express Na+,K+-ATPase, and determine what changes in V-H-ATPase expression occur when freshwater stingrays are acclimated to seawater. The V-H-ATPase is known to play an important role in Na+ uptake and H+ excretion in the gills of freshwater teleosts, and has been identified in gills of a marine elasmobranch. However, V-H-ATPase expression in the gills of a freshwater elasmobranch has never been investigated. Results from this chapter were published in the following paper: “Piermarini, P. M. and Evans, D. H. (2001) Immunochemical analysis of the vacuolar proton-ATPase B-subunit in the gills of a euryhaline stingray (Dasyatis sabina): effects of salinity and relation to Na+/K+-ATPase J Exp Biol 204, 3251-3259”. In Chapter 5, I compare the expression of a Cl-/HCO3exchanger (pendrin) in the gills of freshwater and marine Atlantic stingrays, determine what cells pendrin occurs in, and determine what changes in pendrin expression occur when freshwater stingrays are acclimated to seawater. Pendrin plays an important role in HCO3excretion by the

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21 mammalian kidney, but has not been described in an ion regulatory tissue of a lower vertebrate. Results from this chapter were submitted as a manuscript that is currently being reviewed by the American Journal of Physiology: Regulatory, Integrative, and Comparative Physiology. In Chapter 6, I summarize the results and conclusions presented in this dissertation and propose directions for future research.

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22 Seawater orRectal Gland LumenBlood3 Na+2 K+ ClNa+K+2 Cl-Na+ Teleost Chloride Cell orRectal Gland Tubule CellAccessory CellPavement Cell Seawater orRectal Gland LumenBlood3 Na+2 K+ ClNa+K+2 Cl-Na+ Teleost Chloride Cell orRectal Gland Tubule CellAccessory CellPavement Cell Figure 1-1. Current model of active NaCl secretion in the marine teleost chloride cell and marine elasmobranch rectal gland tubule cell. See text for details. Note that in rectal gland, surrounding cells would be other tubule cells and both tight junctions would be shallow.

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23 Accessory CellPavement CellSeawater Blood3 Na+2 K+ Chloride CellHCO3-ClH+Na+ Accessory CellPavement CellSeawater Blood3 Na+2 K+ Chloride CellHCO3-ClH+Na+ Figure 1-2. Current model of acid/base excretion in the seawater teleost gill. See text for details. For clarity, NaCl secretory pathways are left out.

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24 Fresh waterBlood3 Na+2 K+ Chloride CellHCO3-ClH+Na+ Pavement Cell3 Na+2 K+ H+Na+ Pavement Cell Fresh waterBlood3 Na+2 K+ Chloride CellHCO3-ClH+Na+ Pavement Cell3 Na+2 K+ H+Na+ Pavement Cell Figure 1-3. Current model of active NaCl uptake and acid/base excretion in the freshwater teleost gill. See text for details.

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CHAPTER 2 OSMOREGULATION OF ATLANTIC STINGRAYS FROM THE FRESHWATER ST. JOHNS RIVER, FLORIDA Introduction The common osmoregulatory strategy used by euryhaline elasmobranchs in fresh water is a reduced serum/plasma osmotic pressure, which is primarily attained by a decrease in serum/plasma urea to concentrations that are 30 to 50% lower than those of marine elasmobranchs (Smith 1931, Urist 1962, Thorson et al. 1973). However, euryhaline elasmobranchs in fresh water still maintain relatively high concentrations of urea in their plasma (100 to 250 mmol L-1). The other main organic osmolyte, trimethylamine oxide (TMAO), has not been measured from a euryhaline elasmobranch in fresh water, but based on studies from elasmobranchs in dilute sea water, TMAO is expected to decrease in similar proportion to urea (Goldstein et al. 1968, Goldstein and Forster 1971, Thorson et al. 1973). Serum/plasma concentrations of Na+ and Clare also lowered in freshwater, but not to the same degree as urea (Smith 1931, Urist 1962, Thorson 1967, Thorson et al. 1973). In contrast, stenohaline freshwater stingrays of the family Potamotrygonidae have completely lost the ability to reabsorb urea and TMAO (Thorson et al. 1967, Thorson 1970, Gerst and Thorson 1977), and they maintain plasma ion concentrations much lower than euryhaline elasmobranchs in fresh water (Thorson et al. 1967). Potamotrygonid stingrays can not survive in environments that are hyperosmotic to their plasma (Griffith 25

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26 et al. 1973, Bittner and Lang 1980), and may represent the ultimate stage of elasmobranch evolution in fresh water (Thorson et al. 1983). The size and function of the salt-secreting rectal gland of euryhaline elasmobranchs is reduced in fresh water, because it is not necessary to excrete NaCl in a freshwater environment. Thorson et al. (1978, 1983) reported that freshwater elasmobranchs have smaller rectal gland to body weight (RGBW) ratios than marine species. This has been correlated with a reduction in the number of secretory tubules (Oguri 1964, Thorson et al. 1978) and a lower ATPase activity in the tubule cells (Gerzeli et al. 1976). Stingrays of the family Dasyatidae are well known for their ability to inhabit freshwater environments of Africa, Southeast Asia, Australia, New Guinea, and South America (Thorson and Watson 1975, Taniuchi 1979, Compagno and Roberts 1982, Thorson et al. 1983, Compagno and Roberts 1984, Taniuchi 1991, Compagno and Cook 1995). The only North American stingray species frequently found in fresh water is the Atlantic Stingray, Dasyatis sabina. This coastal, euryhaline species ranges from the Chesapeake Bay to Central America (Bigelow and Schroeder 1953) and is known to enter freshwater rivers on a seasonal basis throughout the Southeastern Atlantic and Gulf of Mexico coasts (Gunter 1938, Sage et al. 1972, Schwartz 1995). Atlantic stingrays also inhabit the St. Johns River of Florida (McLane 1955, Tagatz 1968), where this species has established populations over 300 km from the Atlantic Ocean that reproduce and complete their life cycle in fresh water (Johnson and Snelson 1996). To date, no study has investigated the osmoregulation of Atlantic stingrays in fresh water. De Vlaming and Sage (1973) examined how plasma osmotic parameters of marine Atlantic stingrays responded to diluted concentrations of seawater, and found that

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27 plasma Na+ and Cldecreased to a greater extent than plasma urea in salinities greater than 350 mOsm kg-1 (35% seawater). The lowest salinity that De Vlaming and Sage (1973) exposed marine Atlantic stingrays to was approximately 25% seawater, at which plasma urea concentrations were substantially lower than those from 100% seawater individuals. The goals of this study were to 1) describe the osmoregulation of Atlantic stingrays from the freshwater St. Johns River by measuring plasma osmotic parameters and rectal gland weights; and 2) determine if these freshwater stingrays can osmoregulate in hyperosmotic waters by measuring the same parameters from animals acclimated to higher salinities. I hypothesized that the freshwater populations may show physiological evidence of differentiation from marine Atlantic stingrays (e.g., lack of urea reabsorption), because the St. Johns River Atlantic stingrays reproduce and complete their life cycle in fresh water (Johnson and Snelson 1996). This represents the first study concerning the osmoregulation of Atlantic stingrays in the St. Johns River, and the first experimental determination of how plasma osmolytes of a euryhaline elasmobranch in fresh water respond to a salinity increase. Materials and Methods Field Studies All Atlantic stingrays for this study were collected from Lake Jesup of the St. Johns River, FL using trot-lines baited with shrimp. To measure plasma osmotic parameters of freshwater stingrays from the field, a total of 15 Atlantic stingrays were collected, and immediately after capture, a 0.5 mL blood sample was collected via cardiac puncture using 1 mL heparinized (Na+-heparin) syringe equipped with a 21 or 25 gauge needle. Water samples from Lake Jesup were also collected at this time.

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28 The blood was transferred to 1.5 mL microcentrifuge tubes and transported to Gainesville, FL on ice. The plasma was separated from blood cells with centrifugation (1000 g for 3 min), then transferred to 0.5 mL microcentrifuge tubes and stored at -20C until analyzed. Plasma samples were analyzed for: Na+ and K+ (Radiometer/Copenhagen FLM3 Flame Photometer, Brnshj, Denmark); Cl(Radiometer/Copenhagen CMT10 ClTitrator); Ca2+ (Buck Scientific Model 210 Atomic Absorption Spectrophotometer, East Norwalk, CT); urea (Sigma Blood Urea Nitrogen Kit 535, St. Louis, MO); and total osmolality (Wescor 5100B Vapor Pressure Osmometer, Logan, UT). Water samples were analyzed for the same components, except for urea. Plasma TMAO measurements were attempted using the method of Wekell and Barnett (1991), but values are not reported because of complications with the technique. An additional 32 Atlantic stingrays were collected, then anesthetized with a 0.01% 3-aminobenzoic acid ethyl ester (MS-222, Sigma) solution and pithed. The body weight of these animals was measured, and then rectal glands were dissected out and weighed. RGBW ratios were calculated in mg of rectal gland weight per kg of body weight. These ratios have previously been used as a relative indication of rectal gland function (Thorson et al. 1978, Thorson et al. 1983). Acclimation Experiments Sixteen Atlantic stingrays were collected from Lake Jesup and transported to Gainesville, FL in aerated 120 L coolers. In Gainesville, the stingrays were transferred to 379 L freshwater tanks (four animals per tank). On that same day, blood samples from an additional four stingrays were taken immediately after capture via cardiac puncture, transported back to Gainesville, FL on ice, and the plasma was isolated, stored, and

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29 analyzed as mentioned above. These four animals were not transported to Gainesville and are referred to as the Field group. Eight of the stingrays that were transported to Gainesville, FL remained in fresh water (0.1 parts per thousand; ppt) for the entire experimental period of 32 days, and are referred to as the Control group. The other eight stingrays were left in fresh water for the first 12 days, and then transferred to 50% seawater (15 ppt) over 2 days (approximately 7 ppt per day). After 8 days in 50% seawater the animals were transferred to 100% seawater (30 ppt) over 2 days (approximately 7 ppt per day), where they remained for the last 8 days of the experiment. These stingrays are referred to as the Experimental group. Control and Experimental animals were starved throughout the entire 32-day period to minimize any variation in plasma osmolytes caused by differential feeding of animals. Water temperature in the Control and Experimental tanks ranged from 20 to 23 C; pH was maintained between 7.8 and 8.2 using freshwater and marine pH buffers (Seachem, Stone Mountain, GA). Tanks were equipped with biological filtration to maintain NH3 and NO3 concentrations below 1 part per million. Serial blood samples were taken on Days 12, 22, and 32 via cardiac puncture from Control and Experimental animals. Using a 25 gauge needle attached to a 1 mL heparinized (Na+-heparin) syringe, approximately 0.5 mL of blood was taken from each animal to measure hematocrit and plasma osmolytes. The plasma was separated from blood cells with centrifugation (1000 g for 3 min), then transferred to 0.5 mL microcentrifuge tubes and stored at -20C until analyzed. Water samples were also taken from the holding tanks on each respective day. Both plasma and water samples were analyzed for osmolytes as described above. In addition, on Day 32 all the animals were

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30 anesthetized with a 0.01% MS-222 solution and pithed to obtain their rectal gland and body weights as described above. A potential concern of the blood sampling technique used in this experiment was the effect of stress on plasma osmolytes. Previous studies have shown that plasma Na+, Cl-, and osmolality of some freshwater teleosts was influenced by stress (McDonald and Milligan 1997). In the present study, I did not notice any superficial indicators of stress in the captive animals (e.g., changes in body pigmentation, behavioral modifications). Additionally, a recent study on stress in elasmobranchs found no significant changes in serum Na+ and Clafter exposure to various levels of stress (Manire et al. 2001). Therefore, if the blood sampling caused stress, it is unlikely to affect the measured plasma parameters. Statistical Analyses A one-way repeated measures analysis of variance (ANOVA; P < 0.05) was used to determine if salinity affected plasma osmolality and plasma osmolytes of Experimental stingrays, and if plasma osmolality and plasma osmolytes of the Control group were stable (i.e., equal on Days 12, 22, and 32). A one-way ANOVA (P < 0.05) was used to determine if hematocrit, plasma osmolality, and plasma osmolytes between Control and Experimental groups were different on a given Day. A one-way ANOVA (P < 0.05) was also used to compare RGBW ratios of Control and Experimental animals on Day 32. A two-tailed, unpaired Student’s T-test (P < 0.05) was used to determine if plasma values of the Experimental Atlantic stingrays were equivalent to the values of the tank water on days when blood samples were taken.

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31 Results Plasma Osmolytes and Rectal Glands of Freshwater Atlantic Stingrays Plasma concentrations of Na+, Cl-, urea, K+, and Ca2+ of freshwater Atlantic stingrays were greater than the respective water concentrations from Lake Jesup (Table 2-1). These osmolytes contributed to a plasma osmolality that was 16 times greater than the lake water osmolality (Table 2-1). The RGBW ratio of freshwater Atlantic stingrays was 49.75 2.1 mg kg-1. Acclimation of Freshwater Atlantic Stingrays to Increased Salinity During the acclimation experiment, no mortality occurred in the Control or Experimental groups. Mean hematocrit of Experimental stingrays did not differ significantly from that of Control animals on any day blood samples were taken. The Control and Experimental hematocrit values on Day 32 were 21.5 0.7% and 20.6 0.9%, respectively. Mean plasma osmolality and plasma concentrations of Na+, Cl-, and urea increased significantly in the Experimental stingrays, while these parameters did not change in the Control animals (Figure 2-1). No measured parameter was different between Control and Experimental stingrays on Day 12. On Day 22, Experimental plasma values of osmolality, Na+, Cl-, and urea were 19, 20, 27, and 29% greater than Control plasma values, respectively. On Day 32, Experimental plasma values of osmolality, Na+, Cl-, and urea were 72, 61, 90, and 89% greater than Control plasma values, respectively (Figure 2-1). Plasma concentrations of K+ and Ca2+ increased in the Experimental group and were stable in the Control group, but Experimental groups’ concentrations were not significantly different than the Controls’ until Day 32 (Experimental [K+] = 5.07 0.16

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32 mmol L-1 vs. Control [K+] = 2.99 0.18 mmol L-1; Experimental [Ca2+] = 5.5 0.1 mmol L-1 vs. Control [Ca2+] = 4.62 0.16 mmol L-1). Plasma osmolality and all measured plasma osmolytes in the Experimental stingrays were significantly different than the ambient tank water, except for Na+ on Day 22 (Table 2-2). Plasma of Experimental animals was always hyperosmotic to ambient conditions, except on Day 32 when the plasma was slightly hypoosmotic to ambient water (Table 2-2). Rectal gland weights and RGBW ratios of the Experimental stingrays (34.5 4.0 mg; 60.4 5.4 mg kg-1) were not significantly different than those for the Control group (34.5 2.0 mg; 70.8 3.25 mg kg-1 ). The plasma osmolality and plasma osmolytes (except Ca2+) of Control stingrays on Day 32 were significantly lower than those values in the Field group (Table 2-3). Discussion Osmoregulation of Freshwater Atlantic Stingrays The plasma osmotic composition of freshwater Atlantic stingrays from Lake Jesup is similar to other euryhaline elasmobranchs in fresh water, such as the bull shark, Carcharhinus leucas (Table 2-1). These freshwater stingrays have a 40% lower plasma osmolality than marine Atlantic stingrays, with plasma concentrations of Na+, Cl-, and urea that are approximately 30, 30, and 50% of marine stingray values, respectively (Table 2-1). This is similar to other euryhaline elasmobranchs in fresh water that primarily reduce plasma osmolality through losses of urea, with additional smaller reductions of Na+ and Cl(Smith 1931, Urist 1962, Thorson et al. 1973). Compared to freshwater teleosts and Potamotrygonid stingrays, the freshwater Atlantic stingrays have a substantially higher plasma osmolality and plasma

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33 concentrations of Na+, Cl-, and urea (Table 2-1). The osmotic gradient between the plasma of the freshwater Atlantic stingrays and water of Lake Jesup is almost twice that experienced by a typical freshwater teleost or Potamotrygonid stingray. Therefore, the freshwater Atlantic stingrays may have high urine flow rates to prevent volume loading, as found in other euryhaline elasmobranchs in fresh water (Smith 1931), especially if the freshwater Atlantic stingrays have a similar high permeability to water as found in Potamotrygonid stingrays (Carrier and Evans 1973). Atlantic stingrays in Lake Jesup also maintain large hyperionic gradients of Na+ and Clbetween their plasma and the lake water (Table 2-1), which would result in loss of NaCl to the environment. However, if freshwater Atlantic stingrays have similar low permeability to ions as Potamotrygonid stingrays (Carrier and Evans 1973), the diffusional losses may be small. Although plasma urea concentrations of freshwater Atlantic stingrays are lower than marine individuals, the urea values are still much higher than those found in Potamotrygonid stingrays and freshwater teleosts (Table 2-1). Urea retention does not appear to provide any osmoregulatory advantage, because the freshwater Atlantic stingrays reproduce and complete their life cycle in the St. Johns River. If they excreted more urea, their plasma osmolality would further decrease, which would lead to a lower osmotic gradient and energetic cost of osmoregulation. However, these high basal plasma urea concentrations in freshwater individuals may be a consequence of their kidney anatomy. Although the renal counter current reabsorptive mechanisms of urea in elasmobranchs are still not completely understood, Lacy and Reale (1995) noted a direct correlation between the presence of tubular bundles in nephrons of elasmobranch kidneys

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34 and the renal reabsorption of urea. Potamotrygonid stingrays, who no longer have the capability of reabsorbing urea, do not possess tubular bundles in their kidneys, but kidneys from Atlantic stingrays still have the bundles (Lacy and Reale 1995). It is possible that freshwater Atlantic stingrays and other euryhaline elasmobranchs in freshwater have a limited ability to excrete urea, because of their ancestral renal tubule morphology that still has the mechanisms for urea reabsorption. To balance diffusional and urinary ion losses, it is necessary for freshwater fishes to actively uptake NaCl from the environment with their gills (Perry 1997). Freshwater teleost gills possess mechanisms associated with active ion uptake in their branchial epithelium such as vacuolar H+-ATPase (V-H-ATPase) and Na+,K+-ATPase (Perry 1997, Wilson et al. 2000a). Although marine elasmobranchs have branchial V-H-ATPase and Na+,K+-ATPase (Conley and Mallatt 1988, Wilson et al. 1997), it is not known if these transporters are present in the gills of freshwater species. Further work is needed to determine what transporters are present in the gills of freshwater elasmobranchs and how they may contribute to active NaCl uptake. The RGBW ratios reported in this study for freshwater Atlantic stingrays are 80% lower than values Burger (1972) reported for marine individuals. In other euryhaline elasmobranchs, a smaller rectal gland size has been correlated with a fewer number of secretory tubules and lower ATPase activity (Oguri 1964, Gerzeli et al. 1976), which suggests the glands have a reduced function in fresh water. Therefore, my finding of smaller glands in freshwater Atlantic stingrays may suggest their glands are less active than those from marine individuals. This would be beneficial for ion regulation in a freshwater environment, because NaCl needs to be conserved and not excreted.

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35 Acclimation of Freshwater Atlantic Stingrays to Higher Salinities In this study, hematocrits of Experimental and Control groups were not significantly different from each other, which suggests that freshwater Atlantic stingrays can regulate their blood volume in higher salinities. If they could not regulate blood volume in seawater, I would have expected hematocrits to be significantly higher in the Experimental group, because they would lose plasma water to the ambient seawater. De Vlaming and Sage (1973) suggested that marine Atlantic stingrays regulated their blood volume in dilute seawater, because hematocrits from animals in low salinities were equivalent to marine values after 6 days in a dilute medium. Although plasma osmolality of the Experimental stingrays increased significantly compared to the Controls, the plasma osmolality of Experimental stingrays on Day 32 was almost 100 mOsm kg-1 lower than values reported for marine Atlantic stingrays by De Vlaming and Sage (1973) (Tables 2-1, 2-2). Additionally, the plasma of the Experimental stingrays was hypoosmotic to the ambient 100% seawater (Table 2-2), which is unexpected for an elasmobranch in seawater. Plasma concentrations of Na+ and Clin the Experimental group were not responsible for the osmolality deficit, because they increased to values that were similar to marine Atlantic stingrays (Tables 2-1, 2-2). However, plasma urea concentrations were lower than those found in marine stingrays (Table 2-1, Table 2-2), which may be explained by the effects of starvation on plasma urea (Table 2-3). Experimental and Control groups were starved for 32 days, while marine Atlantic stingrays from De Vlaming and Sage’s (1973) experiments were only starved for 6 days. In the Control stingrays, lower concentrations of plasma Na+, Cl-, and urea were observed on Day 32, compared to Field stingrays (Table 2-3). The Experimental stingrays in

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36 seawater would not lose plasma NaCl due to starvation, because the ambient water is hyperionic to their plasma and NaCl constantly diffuses into the animal. However, urea production is based on the catabolism of free amino acids, which is probably limited during starvation. Additionally, a study by Haywood (1973) found that starvation of marine dogfish (Poroderma africanum) for 32 days induced hypoosmoregulation. This was attributed to a 100 mmol L-1 loss of serum urea; serum Clwas not affected by starvation. Therefore, it is likely that the lower than expected plasma urea and osmolality measured in the Experimental stingrays on Day 32 was caused by the negative effect of starvation on plasma urea concentrations. As previously noted, the plasma Na+ and Clconcentrations of the Experimental group on Day 32 were similar to those reported by De Vlaming and Sage (1973) for marine Atlantic stingrays. This may suggest that rectal glands of Experimental stingrays have a similar ability to excrete NaCl as those of marine individuals. However, the rectal gland weights and RGBW ratios of the Experimental stingrays were not different than those of Control stingrays. This may suggest that the smaller rectal glands of the Experimental group increase their biochemical potential (e.g., Na+, K+-ATPase activity) for NaCl secretion, and an increase in size may not be noticed unless the Experimental animals were exposed to seawater for a longer time period. Another possibility is that the gills and kidneys have a compensatory role in NaCl excretion until the rectal gland reaches its full size. Further biochemical and physiological studies would be required before this can be determined. Evolutionary and Ecological Implications To date, no study has found any major differences between freshwater and marine populations of the Atlantic stingray (Johnson and Snelson 1996, Amesbury and Snelson

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37 1997). This study did not find any major physiological differences between Lake Jesup Atlantic stingrays and marine individuals. Although the St. Johns River stingrays reproduce and complete their life cycle in fresh water (Johnson and Snelson 1996), they have not evolved the osmoregulatory strategy seen in the Potamotrygonid rays. This is probably a result of the Atlantic stingray’s relatively recent occupation (late Pleistocene, approximately 10,000 years) of the St. Johns River (Amesbury and Snelson 1997) and/or lack of appropriate genetic isolation from marine individuals. In contrast, Potamotrygonid stingrays were isolated in the Amazon River Basin for over 65 million years (Thorson et al. 1983). An ecological implication of the results from this study is that the freshwater Atlantic stingrays may enter estuarine and marine environments. Since these animals complete their life cycle in the St. Johns River and can be as far as 300 km from the ocean, it is unlikely that they consistently migrate to seawater. However, the results from the present study suggest that freshwater Atlantic stingrays still have the osmoregulatory mechanisms to survive in higher salinities. Therefore, it is possible that freshwater individuals migrate to the mouth of St. Johns River, similar to the bull sharks of Lake Nicaragua that live in fresh water for extended periods of time, but return to the brackish water mouth of the San Juan River to reproduce (Thorson 1976). An extensive tag and release study on the freshwater Atlantic stingrays would help determine if they are an ecologically euryhaline population.

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38 Table 2-1. Comparison of osmolality and osmolytes in freshwater Atlantic stingray plasma to Lake Jesup water, other freshwater elasmobranchs, a freshwater teleost, and marine Atlantic stingrays. Species Na+ ClUrea K+ Ca2+ Osm D. sabina (FW) a 211.9 2.8 207.8 3.4 195.9 7.9 5.20 0.25 4.3 0.24 621.4 10.8 Lake Jesup Water a 3.0 1.4 3.7 1.5 — 0.07 0.03 1.1 0.16 38.0 0.5 C. leucas (FW) b 245 4.1 219.0 4.9 169.0 5.4 6.40 0.30 4.5 0.15 673.3 12.0 Potamotrygon (FW) c 164.0 5.6 151.7 5.0 1.1 0.1 4.45 0.25 3.0 0.40 282.0 16.8 Teleost (FW) d 130 125 — 2.9 2.1 274 D. sabina (SW) e* 310.0 5.0 300.0 4.5 394.5 5.5 6.95 0.70 3.1 0.20 1034.0 7.5 Note. "Osm" = "Osmolality"; “FW” = “Fresh water”; “SW” = “Seawater” All units are mmol L-1, except for “Osm” which is mOsm kg-1. Values are means 1 S.E (if available). Data from: a This study; b Thorson et al. (1973) and Otake (1991); c Griffith et al. (1973); d Evans (1993); e De Vlaming and Sage (1973) * Values averaged from D. sabina in 98% and 100% SW (Animals starved for 6 days).

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39 Table 2-2. Comparison of osmolality and osmolytes in ambient water and plasma osmolality of Experimental Atlantic stingrays. Na+ ClUrea K+ Ca2+ Osm Ambient (Day 12) 1.25 0.35 1.25 0.04 — <0.1 0.95 0.08 42.17 3.06 Plasma (Day 12) *202.29 5.53 *176.27 2.86 205.08 7.36 *3.44 0.17 *4.79 0.06 *576.14 17.14 Ambient (Day 22) 230.75 2.47 270.88 20.68 — 4.9 0.14 5.93 0.45 496.33 16.97 Plasma (Day 22) 233.25 2.36 *202.66 4.76 227.85 5.87 *3.43 0.18 *4.79 0.05 *652.13 15.43 Ambient (Day 32) 461.00 4.24 565.27 7.16 — 9.83 0.25 11.33 0.80 995.33 0.47 Plasma (Day 32) *319.13 4.20 *296.1 3.69 329.76 4.63 *5.07 0.16 *5.50 0.10 *953.04 14.69 Note. "Osm" = "Osmolality" All units are mmol L-1, except for “Osm” which is mOsm kg-1. Values are means 1 S.E (if available) Asterisks indicate that plasma value was significantly different (P < 0.05) than the ambient water for that Day.

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Table 2-3. Comparison of plasma osmolality and osmolytes in Control group (Day 32) and Field group. 40 Na+ ClUrea K+ Ca2+ Osm Field (N = 4) 220.38 7.65 206.1 9.91 229.01 20.16 4.56 0.48 4.95 0.31 662.08 28.53 Control (N = 8) *198.77 4.15 *156.01 6.86 *174.59 6.05 *2.99 0.18 4.62 0.17 *554.96 14.42 Note. "Osm" = "Osmolality" All units are mmol L-1, except for “Osm” which is mOsm kg-1. Values are means 1 S.E. The Field group represent animals sampled from Lake Jesup on the same day Control animals were transported to the lab. It is assumed that plasma composition of the Field stingrays represents the plasma osmotic makeup of the control group before transportation. Asterisks indicate that the plasma from the Control group was significantly different than the Field group (P < 0.05).

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41 D 100% SW 50% SW FW FW FW FW * * [Urea] (mmol L-1) Time (Day) 32 22 12 400 300 200 100 0 A 100% SW 50% SW FW FW FW FW * * Osmolality (mOsm kg-1) Time (Day) 32 22 12 1000 750 500 250 0 C 100% SW 50% SW FW FW FW FW * * [Cl-] (mmol L-1) Time (Day) 32 22 12 400 300 200 100 0 B 100% SW 50% SW FW FW FW FW * * [Na+] (mmol L-1) Time (Day) 32 22 12 400 300 200 100 0 Figure 2-1. Osmotic composition of plasma from Control (open bars) and Experimental (shaded bars) groups on Days 12, 22, and 32. For the Experimental group, days 12, 22, and 32 represent fresh water, 50% seawater, and 100% seawater, respectively. Values are means + 1 S.E. Asterisks indicate that plasma from the experimental group was significantly different than the control group for that day (P < 0.05).

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CHAPTER 3 EFFECT OF ENVIRONMENTAL SALINITY ON NA+,K+-ATPASE IN THE GILLS AND RECTAL GLAND OF THE ATLANTIC STINGRAY Introduction The teleost gill epithelium has been studied extensively for its role in the acclimation of euryhaline teleost fishes to both freshwater and marine environments. The Na+,K+-ATPase is one of the most important enzymes associated with ion regulation in the fish gill, because it indirectly energizes the branchial excretion of NaCl in marine teleosts (reviewed by McCormick 1995, Marshall and Bryson 1998, Evans et al. 1999). The role of Na+,K+-ATPase in freshwater teleost gills is not as well understood, but it may be important for establishing electrochemical gradients necessary for NaCl and/or calcium uptake (Perry 1997). Na+,K+-ATPase is a membrane bound P-ATPase composed of at least three subunits: , , and (Blanco and Mercer 1998). This ubiquitous transporter is involved with maintaining routine cell volume and resting membrane potential (Blanco and Mercer 1998), and is extremely abundant in specialized ion transporting cells. Two well studied examples of these cells in fishes are the marine teleost chloride cell and elasmobranch rectal gland tubule cell. In these two cell types, Na+,K+-ATPase is localized to the complex basolateral membrane/tubule system (Karnaky et al. 1976, Eveloff et al. 1979) that amplifies the possible sites for Na+,K+-ATPase insertion (Karnaky 1986, Valentich et al. 1995). 42

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43 Many researchers have reported a positive correlation between environmental salinity and biochemical activity of branchial Na+,K+-ATPase in teleost fishes, especially in salmonids and anguillids (Kultz and Jurss 1993, Madsen et al. 1995, McCormick 1995, Uchida et al. 1996, Uchida et al. 1997, Crockett 1999, D'Cotta et al. 2000). However, in other teleost species, branchial Na+,K+-ATPase activity in freshwater-acclimated individuals is equivalent to or exceeds that of seawater-acclimated individuals (Yoshikawa et al. 1993, Madsen et al. 1994, McCormick 1995, Jensen et al. 1998, Vonck et al. 1998, Kelly et al. 1999, Marshall et al. 1999). Recently, further insights into the cellular regulation of fish gill Na+,K+-ATPase were made using heterologous and homologous antibodies and partial cDNA sequences to the -subunit. For example, changes in branchial Na+,K+-ATPase activity were positively correlated with Na+,K+-ATPase-specific mRNA and protein levels (Kisen et al. 1994, Cutler et al. 1995, Madsen et al. 1995, Hwang et al. 1998, Jensen et al. 1998, Lee et al. 1998, D'Cotta et al. 2000). This suggests that changes in fish gill Na+,K+-ATPase activity are due to changes in the relative abundance of the transporter. In mammals, activation of latent Na+,K+-ATPase has also been shown to be responsible for changes in activity of the enzyme (Ewart and Klip 1995), but this has not been studied in fish. Na+,K+-ATPase-rich (chloride) cells have been immunolocalized on both gill lamellae and interlamellar regions of freshwater chum salmon (Oncorhynchus keta), but the cells are only present on the interlamellar region of seawater-acclimated individuals (Ewart and Klip 1995, Uchida et al. 1996, Ura et al. 1996, Uchida et al. 1997, Shikano and Fujio 1998a). It was suggested that Na+,K+-ATPase-rich cells found on the lamellae are specialized for ion uptake from fresh water, whereas interlamellar Na+,K+-ATPase

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44 rich cells are involved with NaCl excretion in a marine environment (Laurent and Dunel 1980, Uchida et al. 1996, Seidelin and Madsen 1999). The guppy (Poecilia reticulata) does not follow the above trend, because Na+,K+-ATPase-rich cells are only found in interlamellar regions of freshwaterand seawater-acclimated fish (Shikano and Fujio 1998c, Shikano and Fujio 1998b, Shikano and Fujio 1999). In contrast to teleost fishes, the role of the gill epithelium in elasmobranch ion regulation has not been well studied. It is thought that the gills do not play a large role in NaCl excretion, because elasmobranchs have a salt-secreting-rectal gland (Shuttleworth 1988, Karnaky 1998). However, a few studies have suggested that the gills are a potential site of net salt secretion, because plasma NaCl concentrations were maintained well below seawater when rectal glands were surgically removed from spiny dogfish, Squalus acanthias (Burger 1965, Evans et al. 1982). In addition, recent evidence of natriuretic peptide receptors in the gills of two marine elasmobranch species (Donald et al. 1997, Sakaguchi and Takei 1998), may indicate the gills are involved with NaCl excretion. In freshwater elasmobranchs, the gills are presumably the site of NaCl uptake, but no studies have examined the branchial epithelium of these fishes. Mitochondrionand Na+,K+-ATPase-rich cells have been described from the gills of a few marine elasmobranch species (Laurent and Dunel 1980, Laurent 1984, Conley and Mallatt 1988). In contrast to chloride cells of marine teleost gills, elasmobranch mitochondrion-rich cells lack a complex basolateral membrane/tubule system, an apical crypt, and accessory cells. Branchial Na+,K+-ATPase activity was measured in marine S. acanthias (Jampol and Epstein 1970, Morgan et al. 1997), and the activity was considerably lower than that found in teleost gills and the elasmobranch rectal gland.

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45 The activity or localization of Na+,K+-ATPase has not been determined in the gills of an elasmobranch acclimated to a salinity lower than 100% seawater. The mechanisms and regulation of NaCl secretion are relatively well established in the marine elasmobranch rectal gland, where Na+,K+-ATPase plays an important role in energizing active NaCl transport (reviewed by Riordan et al. 1994, Valentich et al. 1995, Silva et al. 1997). However, the function of the rectal gland in elasmobranchs from low salinity environments has not been thoroughly investigated. Oguri (1964) and Thorson et al. (1983) showed a positive correlation between relative rectal gland size and environmental salinity that suggested rectal glands from freshwater elasmobranchs had a decreased function, compared to marine individuals. Only Gerzeli et al. (1976) have examined biochemical changes associated with the rectal gland in lower salinities, and they showed that rectal gland tubules from freshwater bull sharks (Carcharhinus leucas) had a lower total ATPase activity than those from marine bull sharks. Although a salt-secreting gland may appear to have no ion regulatory function in a freshwater elasmobranch, the rectal gland is still composed of numerous secretory tubules in both euryhaline and stenohaline freshwater species (Oguri 1964, Thorson et al. 1978). The Atlantic stingray (Dasyatis sabina) is an excellent model species for studying elasmobranch osmoregulatory mechanisms, because it is one of the few elasmobranch species that has established reproducing populations in both freshwater and marine environments. In the previous chapter, I showed that freshwater Atlantic stingrays from the St. Johns River, FL had reduced concentrations of major plasma osmolytes and smaller rectal gland sizes, compared to marine Atlantic stingrays. The freshwater stingrays were capable of acclimating to seawater, where plasma NaCl and urea

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46 concentrations of the acclimated rays were similar to marine stingrays, but their rectal gland size did not appreciably change. The goals of this study were to compare the activity, relative amount, and localization (gills only) of Na+,K+-ATPase in the gills and rectal gland of Atlantic stingrays from freshwater and marine environments. I also compared the same parameters in freshwater stingrays acclimated to seawater for one week. This study is the first to show an effect of environmental salinity on the activity and expression of Na+,K+-ATPase in the gills and rectal gland of an elasmobranch. In addition, this study is the first to immunolocalize Na+,K+-ATPase-rich cells in the gills of an elasmobranch. Materials and Methods Animal Collection and Holding Conditions Ten Atlantic stingrays were captured from the St. Johns River, FL (Lake Jesup or Lake George) with trot lines, transported to Gainesville, FL, and held in two 379 L freshwater, closed-system tanks (five rays per tank; < 1 ppt salinity). In addition, five marine Atlantic stingrays were captured via hook and line from Cedar Key, FL, transported to Gainesville, FL, and held in a 379 L seawater, closed-system tank (32 ppt). Five of the freshwater stingrays were left in fresh water (referred to as freshwater stingrays), while the other five were gradually acclimated to seawater as follows (referred to as seawater-acclimated stingrays). After one week in fresh water, the salinity was raised to 16 ppt over 2 days (8 ppt per day). After two days in 16 ppt, the salinity was raised to 32 ppt seawater over three days. The animals remained in 32 ppt seawater for one week before tissue samples were taken. The marine stingrays from Cedar Key remained in 32 ppt seawater for the entire period (referred to as seawater stingrays). All

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47 animals were fed live grass shrimp (Palaemonetes sp.) every other day, and were starved 48 h before tissue collection. Water temperature of all tanks was maintained at 25C, and pH was adjusted to 8.2 using freshwater and marine buffers (Seachem). The tanks were also equipped with biological filtration, which maintained ambient NH3 and NO3 levels below 1 part per million. Collection of Tissues Animals were anesthetized in 4 L of a 0.01% 3-aminobenzoic acid ethyl ester (MS-222, Sigma) solution made with tank water. For freshwater stingrays, the solution was buffered with a commercial freshwater pH buffer (Seachem) to prevent acidification by the MS-222. Once anesthetized, animals were placed ventral side up in a slanted water bath with their gills immersed in the anesthetic. In order to clear the gills and rectal gland of red blood cells, the animal was perfused with a marine elasmobranch Ringer's solution at 4C (Forster et al. 1972). However, for freshwater stingrays, NaCl, urea, and trimethylamine oxide concentrations in the Ringer’s were reduced to 200, 200, and 41 mmol L-1, respectively. The skin ventral to the heart and pericardium was removed, and 0.5 to 1.0 ml of blood was removed from the ventricle with a heparinized 25 gauge needle attached to a 1 ml syringe. An equal volume of heparinized Ringer's solution was then injected into the ventricle and allowed to circulate for a few minutes. A cannula, connected to a perfusion bottle (1 m above animal), was inserted into the conus arteriosus and held by forceps. Once the perfusion was started, the sinus venosus was cut to relieve backpressure. The perfusion was continued until the gills appeared bleached and the fluid exiting the pericardial cavity was clear of blood (usually 3 to 5 min).

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48 Immediately after the perfusion, the animal was pithed, and the second left and right gill arches were removed and placed in an elasmobranch Ringer's solution on ice. For immunohistochemistry, gill filaments were trimmed off the arches and placed in fixative (3% paraformaldehyde, 0.05% glutaraldehyde, 0.05% picric acid in 10 mmol L-1 phosphate buffered saline, pH 7.3) for 24 h at 4C, then transferred to two changes of 75% ethanol for removal of fixative. Tissues were left in the second change of 75% ethanol until embedded. Additional filaments were 1) snap frozen in liquid nitrogen for immunoblot analysis, and 2) placed in an ice cold buffer containing 250 mmol L-1 sucrose, 10 mmol L-1 Na2EDTA, and 50 mmol L-1 imidazole (SEI buffer; pH 7.3) that was then frozen on dry ice for Na+,K+-ATPase activity measurements. The rectal gland was then excised and pieces were taken for immunoblot and Na+,K+-ATPase activity measurements. All samples for immunoblots and enzyme assays were stored at -80C. Na+,K+-ATPase Activity Activity of Na+,K+-ATPase was measured using a technique developed by McCormick (1993). Frozen tissue in SEI buffer was immediately thawed and homogenized, and the homogenate was centrifuged at 5000 x g for 30 s at 4C. A 10 l sample of supernatant was added to 2 wells of a 96 well microplate on ice, then 200 l of an assay mixture (50 mmol L-1 imidazole, 45 mmol L-1 NaCl, 10 mmol L-1 KCl, 5 mmol L-1 MgCl2, 2.8 mmol L-1 phosphoenolpyruvate, 0.22 mmol L-1 NADH, 0.7 mmol L-1 adenosine triphosphate, 4.6 U mL-1 lactate dehydrogenase, and 5.1 U mL-1 pyruvate kinase) with or without ouabain (0.7 mmol L-1) was added to a well. The Na+,K+-ATPase activity was determined by subtracting the oxidation rate of NADH to NAD (at 340 nm)

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49 in the presence of ouabain from the oxidation rate without ouabain, at 25 C. Absorbance was measured using a Spectramax 250 microplate reader (Molecular Devices, Sunnyvale, CA). Total protein of the supernatant was also measured using a bicinchoninic acid (BCA) assay (Pierce, Rockford, IL). Activity measurements were expressed as mol of ADP mg protein-1 h-1. Immunoblots Preparation of tissue for immunoblots was similar to Claiborne et al. (1999), with modifications. On a single day, gill filaments or a piece of rectal gland from a freshwater, seawater-acclimated, and seawater stingray were prepared. First, the tissue was placed in ice cold homogenization buffer (250 mmol L-1 sucrose, 1 mmol L-1 Na-EDTA, 2 g mL-1 aprotinin, 2 g mL-1 leupeptin, 100 g mL-1 phenyl methylsulfonyl fluoride and 30 mmol L-1 Tris), and was homogenized with a Tissue-tearor (Biospec Products, Bartlesville, OK) in a 4C cold room on ice. Homogenates were filtered through cheese cloth and centrifuged (3000 g) for 5 min at 4C to remove nuclei and debris. The supernatant was then filtered through cheesecloth and centrifuged (50,000 g) for 30 min at 4C to pellet membrane fractions. The pellet was resuspended with a minimal volume of ice cold homogenization buffer, then an equal volume of a modified Laemmli sample buffer (Laemmli 1970), without bromophenol blue and -mercaptoethanol, was added to solubilize the proteins. The resulting protein samples were centrifuged for 5 to 10 s at 16,000 g to pellet any undissolved material. The total protein content of the supernatant was determined with a detergent compatible assay (Bio-Rad, Hercules, CA), after which bromophenol

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50 blue and -mercaptoethanol were added to final concentrations of 0.01% and 2%, respectively. A 20 g sample of total gill membrane protein (5 g for rectal gland membrane protein) from one stingray in each condition was loaded in triplicate and run on a 7.5% Tris-HCl precast polyacrylamide gel (Bio-Rad) for 1 h at 125 V. Note that it was necessary to load four times more total protein from gill samples, because gills contained a lower abundance of the -subunit per g total membrane protein, compared to the rectal gland. Proteins were then transferred to a polyvinylidene difluoride membrane (PVDF; Bio-Rad) using a wet (20% MeOH, Tris-glycine) transfer unit for 2.5 h at 90 volts in a 4C cold room with stirring. The protein preparation, electrophoresis, and blotting were repeated for the remaining gill and rectal gland samples, which resulted in a total of five PVDF membranes for each tissue with each containing, in triplicate, gill or rectal gland membrane protein from a single freshwater, seawater-acclimated, and seawater stingray. Each PVDF membrane was blocked with blocking buffer (Tris-buffered saline with 5% non-fat dry milk, 0.1% Tween-20, and 0.02% NaN3, pH 7.4) for 1.5 h at 25C, and then transferred to an antibody solution (monoclonal antibody “a5” culture supernatant diluted 1:1000 in blocking buffer) overnight at 4C. The primary antibody, “a5”, developed by Dr. Douglas Fambrough was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA 52242. This antibody was raised against the -subunit of avian Na+,K+-ATPase, and has been used several times to study Na+,K+

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51 ATPase in fish gills (Witters et al. 1996, Choe et al. 1999, Schreiber and Specker 1999, Dang et al. 2000b). After antibody incubation, the PVDF was washed three times (15 min each) with Tris-buffered saline containing 0.1% Tween-20 (TTBS, pH 7.4), then incubated with an alkaline-phosphatase conjugated goat anti-mouse IgG secondary antibody (Bio-Rad; diluted 1:3000 in blocking buffer) for 2 h at 25C. The PVDF was then washed three times (15 min each) with TTBS, and a substrate solution (Bio-Rad Immun-Star ECL Kit) was applied to the PVDF for 5 min at 25 C to initiate a luminescent signal. Binding of antibody was detected by exposing Hyperfilm-ECL imaging film (Amersham, Piscataway, NJ) to the PVDF membrane. Negatives were digitized into TIFF files using a flatbed scanner with transparency adapter (UMAX, Dallas, TX). As a control, PVDF membranes were incubated with normal goat serum diluted in blocking buffer, instead of the anti-Na+,K+-ATPase monoclonal antibody. To quantify the relative abundance of Na+,K+-ATPase immunoreactivity, I measured the optical density (uncalibrated) of the immunopositive band in each animal using Scion Image version 4.02 (Scion Corporation, Frederick, MD). On a given blot, optical density values were measured for each animal, and then standardized to the freshwater condition to calculate relative abundance. Therefore, all relative abundance measurements of freshwater gills were “1.0”, and those of seawater-acclimated and seawater gills were a fraction of the freshwater value. Immunohistochemistry The fixed gill tissue (stored in 75% ethanol) was dehydrated in an ethanol series, and embedded into paraffin wax. Serial sections of gill tissue, parallel to the long axis of the filament, were cut at 6 m and placed on poly-L-lysine coated slides (three sections

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52 per slide). Longitudinal sections of gill filaments provide cross-sectional orientations of gill lamellae, which appear as finger-like projections, and interlamellar regions, which are basal to and between lamellae (see Figure 3-4). For immunostaining, only slides containing sections from the trailing (afferent) half of the gill lamellae and filament were chosen, because preliminary trials showed that Na+,K+-ATPase-rich cells were found exclusively on the trailing half of gill lamellae and filaments (unpublished observation). This is similar to what has been reported for teleost chloride cells (Van Der Heijden et al. 1997). The sections were deparaffinized in Hemo-De (Fisher Scientific, Pittsburgh, PA), hydrated in an ethanol series, and washed in 10 mmol L-1 phosphate buffered saline (PBS). A hydrophobic PAP-Pen (Electron Microscopy Sciences, Fort Washington, PA) was used to draw circles around the tissue sections, and then 3% H2O2 was placed on the sections for 30 min to inhibit endogenous peroxidase activity. Sections were also blocked with a casein solution (Powerblock, Biogenex, San Ramon, CA) for 5 min. The monoclonal antibody “a5” (diluted 1:175 in PBS) was incubated on the sections for 2 h at 25C. The antibody was rinsed off with PBS and the sections were soaked in PBS for 5 min. The sections were then incubated with a biotinylated goat anti-mouse IgG secondary antibody (Biogenex) and a horseradish peroxidase-labeled strepavidin solution (Biogenex) for 20 min each at 25C (with a 5 min PBS wash after each incubation). Antibody binding was visualized by applying the chromagen 3,3’-diaminobenzidine tetra-hydrochloride (DAB; Biogenex) to the sections for 5 min at 25C. In each experiment, one section was exposed to PBS in place of the anti-Na+,K+-ATPase monoclonal antibody as a negative control.

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53 Quantification of the immunohistochemistry was performed by counting the number of immunopositive (Na+,K+-ATPase-rich) cells per gill lamella and per interlamellar space. For each animal, three immunostained slides were chosen randomly. On a section from each slide, the number of Na+,K+-ATPase-rich cells was counted on 30 randomly selected lamellae and interlamellar regions. Lengths of lamellae were also measured to standardize lamellar cell counts. Results are expressed as number of Na+,K+-ATPase-rich cells per 100 m of lamella, per interlamellar region, and per 100 m of lamella + interlamellar region (referred to as ‘sum’). Statistical Analyses For mean Na+,K+-ATPase activities and number of Na+,K+-ATPase-rich cells, differences among groups were detected using an one-way ANOVA, with a Student-Newman-Keuls multiple comparisons test. Differences in relative band intensities from immunoblots were detected using a Kruskal-Wallis non-parametric ANOVA, with a Kruskal-Wallis multiple comparisons test (Conover 1980). All tests were 2-tailed and differences were considered significant if P < 0.05. Results Na+,K+-ATPase Activity In gills, Na+,K+-ATPase activity was highest in freshwater, intermediate in seawater-acclimated, and lowest in seawater Atlantic stingrays (Figure 3-1 A). In rectal glands, Na+,K+-ATPase activity was lowest in freshwater stingrays and highest in seawater-acclimated and seawater stingrays, whose activities were statistically equivalent (Figure 3-1 B).

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54 Immunoblots The anti-Na+,K+-ATPase -subunit monoclonal antibody bound to a protein of 110.9 1.0 kDa in gills and 111.7 1.4 kDa in rectal glands (Figure 3-2). No detectable signal was present when PVDF membranes were incubated with normal goat serum instead of the primary antibody (data not shown). In the gills, relative abundance of the -subunit immunoreactivity followed a similar trend as Na+,K+-ATPase activities; -subunit immunoreactivity was highest in freshwater, intermediate in seawater-acclimated, and lowest in seawater stingrays (Figure 3-3 A). In the rectal gland, relative abundance of the -subunit immunoreactivity also followed a similar trend as Na+,K+-ATPase activities; -subunit immunoreactivity was lowest in freshwater and highest in seawater-acclimated and seawater stingrays, whose relative abundances were statistically equivalent (Figure 3-3 B). Immunohistochemistry Immunostaining for Na+,K+-ATPase was restricted to the basolateral region of relatively large cells in the gill epithelium, and not found on pavement cells (Figure 3-4). No staining was detected when PBS was used instead of the primary antibody (data not shown). The number of Na+,K+-ATPase-rich cells per 100 m of lamella and “sum” number (per 100 m of lamella + per interlamellar region) of Na+,K+-ATPase-rich cells followed the same trend as Na+,K+-ATPase activity and immunoreactivity; freshwater stingrays had the most, seawater-acclimated stingrays had intermediate, and seawater stingrays had the least (Figure 3-5 A, C). In the interlamellar region, number of Na+,K+-ATPase-rich cells was highest in gills of seawater-acclimated stingrays, intermediate in freshwater, and lowest in seawater individuals (Figure 3-5 B).

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55 Additionally, there appeared to be qualitative trends in the intensity of Na+,K+-ATPase immunostaining. In freshwater stingrays, staining appeared to be darkest in Na+,K+-ATPase-rich cells on the distal parts of lamellae, and was weakest in cells on the basal parts of lamellae and the interlamellar region (Figure 3-4 A). In seawater-acclimated stingrays, staining in Na+,K+-ATPase-rich cells was dark on both the lamellae and interlamellar regions (Figure 3-4 B). In seawater stingrays, staining in the Na+,K+-ATPase-rich cells was moderate in the interlamellar region (Figure 3-4 C). Discussion Gills The results from this study are the first to show that activity and expression of Na+,K+-ATPase in the gills of a euryhaline elasmobranch are influenced by environmental salinity. Branchial Na+,K+-ATPase activity was highest in freshwater stingrays, followed by seawater-acclimated, and seawater animals (Figure 3-1 A). In salmonid and anguillid teleosts, gill Na+,K+-ATPase activity is positively correlated with environmental salinity, but in other fishes, a negative or no correlation with salinity is often found (McCormick 1995, Jensen et al. 1998, Kelly et al. 1999, Marshall et al. 1999). Immunoblotting with an anti-Na+,K+-ATPase -subunit antibody showed the presence of an approximate 111 kDa protein in the gill membrane enrichments (Figure 3-2), which is the expected size for the -subunit in vertebrates (Blanco and Mercer 1998) Semi-quantitative immunoblotting revealed that the relative abundance of Na+,K+-ATPase immunoreactivity was influenced by salinity (Figure 3-3 A), in a similar manner as the activity measurements. This suggests that salinity-associated changes of Na+,K+

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56 ATPase activity in the Atlantic stingray gill are accomplished by changing relative amounts of the Na+,K+-ATPase protein, possibly through the number of Na+,K+-ATPase-rich cells on gill lamellae (Figure 3-5 A). If elasmobranch gill Na+,K+-ATPase activity is regulated in a similar manner to teleosts, then the lower Na+,K+-ATPase -subunit immunoreactivity found in seawater-acclimated and seawater stingrays may be a result of a decrease in -subunit mRNA levels (Madsen et al. 1995, Hwang et al. 1998, Jensen et al. 1998, Cutler et al. 2000, D'Cotta et al. 2000), through a decrease in transcription or an increase in mRNA degradation. In salmonids, the turnover rate of gill Na+,K+-ATPase protein does not appear to be affected by salinity (D'Cotta et al. 2000). The Na+,K+-ATPase activity measurements reported in this study are the first for gills of a freshwater elasmobranch. The higher branchial Na+,K+-ATPase activity associated with freshwater stingrays (compared to seawater individuals) may indicate that the gills have an increased active transport role in fresh water, possibly for the absorption of NaCl. An alternative explanation is that the higher Na+,K+-ATPase activity is associated with an increased cost of generic cell volume regulation in all freshwater gill cells, but this is unlikely, because Na+,K+-ATPase immunostaining occurred in specific cell types and not across the entire gill epithelium (Figure 3-4). The lower branchial Na+,K+-ATPase activity measurements in the seawater-acclimated and seawater stingrays was expected, because elasmobranchs have a salt-secreting rectal gland that is the primary site of NaCl excretion (Shuttleworth 1988). Although the branchial Na+,K+-ATPase activity is relatively low in seawater-acclimated and seawater stingrays, it may be enough to energize some active NaCl excretion,

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57 especially in situations where the rectal gland is removed (Evans et al. 1982) or reduced in size (see below). Na+,K+-ATPase immunostaining occurred in relatively large cells of the gill epithelium that are likely mitochondrion-rich, because an abundance of Na+,K+-ATPase has been shown to be a reliable marker of mitochondrion-rich cells in gills of other fishes (Uchida et al. 1996, Witters et al. 1996), including hagfish (Choe et al. 1999). The Na+,K+-ATPase-rich cells were most common and stained the darkest on lamellae of freshwater stingrays, which suggests they might be involved with active ion uptake. In freshwater-acclimated teleost gills, Na+,K+-ATPase-rich (chloride) cells were reported on lamellae (Avella et al. 1987, Uchida et al. 1996, Ura et al. 1996, Seidelin and Madsen 1999, Wong and Chan 1999), and were hypothesized to be sites of ion uptake from the freshwater environment (Laurent and Dunel 1980, Avella et al. 1987, Uchida et al. 1996). However, processes of ion uptake and acid-base excretion are inextricably linked in the gills of freshwater fish (Claiborne 1998, Evans et al. 1999). Therefore, the Na+,K+-ATPase-rich cells may also be involved with the excretion of acid (H+) and/or base (HCO3-). The greater number of interlamellar Na+,K+-ATPase-rich cells in seawater-acclimated stingrays, and the darker immunostaining of interlamellar Na+,K+-ATPase-rich cells in seawater-acclimated and seawater stingrays, may suggest that these cells are involved with active NaCl excretion. In seawater-acclimated teleosts, Na+,K+-ATPase-rich cells in interlamellar regions are thought to be sites of active NaCl excretion (Uchida et al. 1996). However, presence of Na+,K+-ATPase-rich cells in the gills does not necessarily imply active NaCl excretion. Hagfish (Myxine glutinosa) also possess

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58 branchial Na+,K+-ATPase-rich cells (Choe et al. 1999), but are strict ion conformers. Choe et al. (1999) suggested that hagfish Na+,K+-ATPase-rich cells may facilitate acid/base exchanges (Evans 1984a) and/or Ca2+ transport (Forster and Fenwick 1994) known to occur across the hagfish gill epithelium. These functions should also be considered for Na+,K+-ATPase-rich cells found in the gills of seawater-acclimated and seawater stingrays. Rectal Gland The results from this study are also the first to show that activity and expression of Na+,K+-ATPase in the rectal gland of a euryhaline elasmobranch are influenced by environmental salinity. Activity of Na+,K+-ATPase in rectal glands from seawater-acclimated and seawater stingrays was greater than that from freshwater stingrays (Figure 3-1 B). Immunoblotting with an anti-Na+,K+-ATPase -subunit antibody detected an approximate 112 kDa protein in the gill membrane enrichments (Figure 3-2), which is the expected size for the -subunit in vertebrates (Blanco and Mercer 1998). Semi-quantitative immunoblotting revealed that the relative abundance of Na+,K+-ATPase immunoreactivity was influenced by salinity (Figure 3-3 B), in a similar manner as the activity measurements. This suggests that salinity-associated changes of Na+,K+-ATPase activity in the Atlantic stingray rectal gland are accomplished by changing relative amounts of the Na+,K+-ATPase protein. Rectal gland Na+,K+-ATPase activity measurements reported in this study are the first for a freshwater elasmobranch. Although freshwater rectal gland Na+,K+-ATPase activity was about 50% less than that of seawater-acclimated and seawater stingrays, the activity is still about seven times higher than that found in the gills. It is possible that

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59 freshwater stingrays cannot completely turn off active NaCl secretion in the rectal gland, which may be indicative of their relatively recent (in evolutionary time) occurrence in fresh water (see Chapter 2, Amesbury and Snelson 1997). Maintaining a relatively high Na+,K+-ATPase activity in a salt-secreting tissue during exposure to a low salinity may be adaptive for a species that frequently encounters variations in environmental salinity (Marshall et al. 1999). An alternative explanation is that Na+,K+-ATPase activity in the freshwater rectal gland may be related to a secondary function of the gland, such as the active secretion of xenobiotics, which has recently been described in the rectal gland of S. acanthias (Miller et al. 1998). Further biochemical studies on the freshwater stingray rectal gland should help elucidate its function. It was not surprising that Na+,K+-ATPase activity and immunoreactivity were greatest in rectal glands from seawater-acclimated and seawater stingrays, because the rectal gland is the primary site of NaCl excretion in elasmobranchs. However, it is important to note that the size of this gland in seawater-acclimated stingrays is still approximately 50-70% smaller than a typical marine stingray rectal gland (see Chapter 2). Therefore, the total rectal gland Na+,K+-ATPase (rectal gland mass x Na+,K+-ATPase activity) of a seawater-acclimated animal is probably less than that of a seawater stingray, although it may be similar on a “per g protein” level. This may suggest that rectal glands from seawater-acclimated stingrays have a lesser ability to secrete NaCl than those from seawater individuals, and that seawater-acclimated stingrays require extra-rectal gland mechanisms of NaCl excretion, until the gland reaches its full size. A potential site for this NaCl excretion would be branchial Na+,K+-ATPase-rich cells in the

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60 interlamellar region that increased in number and immunostaining during acclimation to seawater (Figures 3-4, 3-5 B). An alternative explanation is that rectal glands of seawater-acclimated stingrays compensate for the reduced mass by increasing the secretion rate and/or NaCl concentration of the secreted fluid, or by increasing the relative abundance of the cystic-fibrosis transmembrane regulator (CFTR) Clchannel, which is thought to be the rate-limiting step in active NaCl excretion (Riordan et al. 1994). Further studies on other transporters involved with active NaCl excretion in the rectal gland (e.g., CFTR, Na+K+2Clcotransporter) as well as functional studies on isolated, perfused glands would be helpful in comparing the relative salt-secreting capabilities of rectal glands from seawater-acclimated and seawater stingrays. In summary, I have presented data that suggest Na+,K+-ATPase activity and expression in the gills and rectal gland of the Atlantic stingray are influenced by environmental salinity. The Na+,K+-ATPase activity and immunoreactivity in gills were negatively correlated with salinity, which suggests the gills might be important for active ion uptake from low salinity environments. In contrast, Na+,K+-ATPase activity and immunoreactivity in the rectal gland were positively correlated with salinity, which supports the hypothesis that the rectal gland is the primary site of NaCl excretion in marine elasmobranchs. Studies on other important ion regulatory transporters (e.g., V-H-ATPase, Na+K+2Clcotransporter, CFTR) in the gills and/or rectal gland would be useful to further describe the rare phenomenon of elasmobranch euryhalinity on a biochemical and cellular level.

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61 a b b a c b B A 1.4 1.2 1.0 0.8 0.6 0.4 0.2 0.0 16 14 12 10 8 6 4 2 0 Na+,K+-ATPase Activity (mol ADP mg protein-1 hr-1) Na+,K+-ATPase Activity (mol ADP mg protein-1 hr-1) FW SWA SW Figure 3-1. Activity of Na+,K+-ATPase in the gills (A) and rectal gland (B) of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) Atlantic stingrays. N = 5 for all groups, except for rectal glands of SWA and SW stingrays where N = 4. Values are means + 1 S.E. Lower case letters above error bars indicate statistical categorization of the means, as determined by a Student-Newman-Keuls multiple comparisons test (P < 0.05).

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62 FW SWA SW Rectal Gland SW SWA Gill FW kDa 43 77 130 205 Figure 3-2. Representative immunoblot for Na+,K+-ATPase in gill and rectal gland membrane enrichments of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) stingrays. Antibody was specific for a 110.9 1.0 kDa protein in gills and a 111.7 1.4 kDa protein in the rectal gland, representing the -subunit of Na+,K+-ATPase (arrow). Black lines represent migration of molecular weight markers (kDa). Note that intensity of gill and rectal gland immunoreactivity cannot be compared to one another, because different amounts of total protein were loaded (20 g for gill, 5 g for rectal gland).

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63 1.2 c b a A 1.0 Relative Abundance 0.8 0.6 0.4 0.2 0.0 4.0 b b a B FW SWA SW 3.5 3.0 Relative Abundance 2.5 2.0 1.5 1.0 0.5 0.0 Figure 3-3. Relative abundance of immunoreactivity for 111 or 112 kDa band representing the -subunit of Na+,K+-ATPase in gills (A) and rectal gland (B) of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) stingrays. N = 5 for all groups. Values are presented as means + 1 S.E. Lower case letters above error bars indicate statistical categorization of groups as determined by a Kruskal-Wallis multiple comparisons test (P < 0.05). Note that no error bars are present in FW, because of the standardization procedure (see Materials and Methods).

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64 A B C Figure 3-4. Representative photomicrographs of Na+,K+-ATPase immunostaining in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). The Na+,K+-ATPase-rich cells (brown) occurred on lamellae (finger-like projections) and/or interlamellar regions (basal to and between lamellae). Bar = 100 m.

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65 a 8 6 4 2 0 C B A c b c b a 5 4 3 2 1 0 c b a 12 10 8 6 4 2 0 Number of Cells (per 100 m of lamella) Number of Cells (per Interlamellar Region) Number of Cells (Sum) SW SWA FW Figure 3-5. Number of Na+,K+-ATPase-rich cells in the gills of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) stingrays. Note that “Sum” = number of Na+,K+-ATPase-rich cells per 100 m of lamella + per interlamellar region. Values are means + 1 S.E. Lower case letters above error bars indicate statistical categorization of the means as determined by a Student-Newman-Keuls multiple comparisons test (P < 0.05).

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CHAPTER 4 IMMUNOCHEMICAL ANALYSIS OF THE VACUOLAR PROTON-ATPASE B-SUBUNIT IN THE GILLS OF THE ATLANTIC STINGRAY: INFLUENCE OF SALINITY AND RELATION TO NA+,K+-ATPASE Introduction The vacuolar-proton-ATPase (V-H-ATPase) is a multi-subunit transporter that is important for energizing a variety of active transport processes in animals (reviewed by Nelson and Harvey 1999). The V-H-ATPase is composed of a catalytic V1 domain that binds ATP and a membrane-bound V0 domain that forms a channel for H+ to cross the cell or vacuolar membrane (Wieczorek et al. 2000). The B-subunit of V-H-ATPase is a component of the V1 domain, and the B-subunit amino acid sequence is very conserved across a wide range of animal taxa (Sudhof et al. 1989, Gill and Ross 1991, Filippova et al. 1998, Niederstatter and Pelster 2000, Perry et al. 2000). Two isoforms of the B-subunit were identified in mammals and fish; using mammalian nomenclature, they are a ubiquitous 58 kDa brain (B2) isoform and a 56 kDa renal (B1) isoform (Nelson et al. 1992, Niederstatter and Pelster 2000). V-H-ATPase was studied extensively in the mammalian renal collecting duct and turtle urinary bladder where immunocytochemical and ultrastructural research has shown that two populations of mitochondrion-rich intercalated cells exist that acidify or alkalinize the urine. Type A (acidifying) intercalated or mitochondrion-rich cells are characterized by apical cell membrane localization of V-H-ATPase, while Type B (alkalinizing) intercalated or mitochondrion-rich cells express V-H-ATPase diffusely throughout the cytoplasm and on their basolateral membrane (Stetson and Steinmetz 66

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67 1985, Brown et al. 1988a, Brown et al. 1988b, Verlander et al. 1992, Verlander et al. 1994b, Brown and Breton 1996, Steinmetz et al. 1996, Brown and Breton 2000). In aquatic vertebrates, V-H-ATPase has been implicated in the energization of NaCl uptake. For example, in amphibian skin it is well established that an apical V-H-ATPase, localized to mitochondrion-rich cells, generates a membrane potential (inside negative) that drives Na+ entry into the epithelium via an apical Na+ channel (Harvey 1992, Ehrenfeld and Klein 1997), and can also energize active Cluptake through apical Cl-/HCO3exchangers (Larsen et al. 1996). Recently, the V-H-ATPase has been considered important for driving Na+ uptake across the gill epithelium of freshwater fishes. Similar to amphibian skin, it was proposed that an apical V-H-ATPase would generate a favorable electrical gradient to drive Na+ uptake through an apical Na+ channel (Lin and Randall 1993, Lin et al. 1994, Sullivan et al. 1995). This model of ion uptake in freshwater teleosts has been supported by the results of Wilson et al. (2000a) who showed that the apical region of pavement cells from tilapia gills (Oreochromis mossambicus) contained immunolabeling for V-H-ATPase and an epithelial Na+ channel (ENaC). In rainbow trout (Oncorhynchus mykiss), apical V-H-ATPase and ENaC were expressed in both pavement and chloride (Na+,K+-ATPase-rich) cells (Wilson et al. 2000a). Other supporting evidence for the proposed role of V-H-ATPase in freshwater teleost ion uptake are bafilomycin inhibition of Na+ uptake in tilapia (O. mossambicus) and carp (Cyprinus carpio) (Fenwick et al. 1999), and decreased activity and immunoreactivity of branchial V-H-ATPase when freshwater rainbow trout are acclimated to seawater (Lin and Randall 1993, Lin et al. 1994).

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68 V-H-ATPase has also been hypothesized to play an important role in systemic acid/base balance of freshwater teleosts, because an apical V-H-ATPase would directly pump H+ into the environment. Experiments on rainbow trout have corroborated this hypothesis by showing that gill V-H-ATPase activity, immunoreactivity, and mRNA expression, all increase after exposure to environmental hypercapnia (Lin and Randall 1993, Lin et al. 1994, Sullivan et al. 1995, Sullivan et al. 1996, Perry et al. 2000). Studies on branchial V-H-ATPase in a true marine teleost have not been published, but the transporter’s role in acid/base regulation of seawater teleosts is assumed to be minimal, because of the favorable gradient for Na+ entry that could drive passive Na+/H+ exchangers (NHEs) (Claiborne 1998, Claiborne et al. 1999). V-H-ATPase activity and immunolabeling were detected in the gills of two marine elasmobranch species, Squalus acanthias and Raja erinacea (Kormanik et al. 1997, Wilson et al. 1997). In gills of S. acanthias, Wilson et al.(1997) found V-H-ATPase immunoreactivity within cytoplasmic vesicles of mitochondrion-rich cells on the interlamellar region. Interestingly, I found that Na+,K+-ATPase immunoreactivity was localized to cells of the interlamellar region in gills from marine Atlantic stingrays (Dasyatis sabina) (see Chapter 3). Therefore, it is possible that V-H-ATPase and Na+,K+-ATPase are expressed in the same branchial cell type of marine elasmobranchs. The goals of this study were to determine if environmental salinity influences expression of V-H-ATPase in the gills of the Atlantic stingray, and determine if V-H-ATPase and Na+,K+-ATPase were expressed in the same cells. To date, studies on the effects of salinity on V-H-ATPase expression in the gills of an elasmobranch have not been published. In Chapter 3, I found that activity and expression of Na+,K+-ATPase in

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69 the Atlantic stingray was 3 to 4 times greater in the gills of freshwater stingrays, compared to marine individuals. I hypothesize a similar trend for V-H-ATPase, because this transporter is considered important for ion uptake in freshwater vertebrates. Materials and Methods Animal Collection and Holding Conditions Ten Atlantic stingrays were captured from the St. Johns River, FL (Lake Jesup or Lake George) with trot lines, transported to Gainesville, FL, and held in two 379 L freshwater, closed-system tanks (five rays per tank; < 1 ppt salinity). In addition, five marine Atlantic stingrays were captured via hook and line from Cedar Key, FL, transported to Gainesville, FL, and held in a 379 L seawater, closed-system tank (32 ppt). Five of the freshwater stingrays were left in fresh water (referred to as freshwater stingrays), while the other five were gradually acclimated to seawater as follows (referred to as seawater-acclimated stingrays). After one week in fresh water, the salinity was raised to 16 ppt over 2 days (8 ppt per day). After two days in 16 ppt, the salinity was raised to 32 ppt seawater over three days. The animals remained in 32 ppt seawater for one week before tissue samples were taken. The marine stingrays from Cedar Key remained in 32 ppt seawater for the entire period (referred to as seawater stingrays). All animals were fed live grass shrimp (Palaemonetes sp.) every other day, and were starved 48 h before tissue collection. Water temperature of all tanks was maintained at 25C, and pH was adjusted to 8.2 using freshwater and marine buffers (Seachem). The tanks were also equipped with biological filtration, which maintained ambient NH3 and NO3 levels below 1 part per million.

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70 Collection of Gill Tissue Animals were anesthetized in 4 L of a 0.01% 3-aminobenzoic acid ethyl ester (MS-222, Sigma) solution made with tank water. For freshwater stingrays, the solution was buffered with a commercial freshwater pH buffer (Seachem) to prevent acidification by the MS-222. Once anesthetized, animals were placed ventral side up in a slanted water bath with their gills immersed in the anesthetic. In order to clear the gills and rectal gland of red blood cells, the animals were perfused with a marine elasmobranch Ringer's solution at 4C (Forster et al. 1972). However, for freshwater stingrays, NaCl, urea, and trimethylamine oxide concentrations in the Ringer’s were reduced to 200, 200, and 41 mmol L-1, respectively. The skin ventral to the heart and pericardium was removed, and 0.5 to 1.0 ml of blood was removed from the ventricle with a heparinized 25 gauge needle attached to a 1 ml syringe. An equal volume of heparinized Ringer's solution was then injected into the ventricle and allowed to circulate for a few minutes. A cannula, connected to a perfusion bottle (1 m above animal), was inserted into the conus arteriosus and held by forceps. Once the perfusion was started, the sinus venosus was cut to relieve backpressure. The perfusion was continued until the gills appeared bleached and the fluid exiting the pericardial cavity was clear of blood (usually 3 to 5 min). Immediately after the perfusion, the animal was pithed, and the second left and right gill arches were removed and placed in an elasmobranch Ringer's solution on ice. For immunohistochemistry, gill filaments were trimmed off the arches and placed in fixative (3% paraformaldehyde, 0.05% glutaraldehyde, 0.05% picric acid in 10 mmol L-1 phosphate buffered saline, pH 7.3) for 24 h at 4C, then transferred to two changes of 75% ethanol for removal of fixative. Tissues were left in the second change of 75%

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71 ethanol until embedded. Additional filaments were snap frozen in liquid nitrogen for immunoblot analysis and stored at -80C until analyzed. Antibodies The antibody to detect V-H-ATPase was developed by Filippova et al.(1998) and is a rabbit polyclonal raised against a 279 amino acid region (amino acids 79 to 357) of the V-H-ATPase B-subunit from the insect Culex quinquefasciatus. This region of the insect B-subunit V-H-ATPase shares 91% amino acid identity with that published for teleost V-H-ATPase B-subunits (Niederstatter and Pelster 2000, Perry et al. 2000). The antibody was kindly provided by Dr. William Harvey, Whitney Laboratory, University of Florida (with permission from Dr. Sarjeet Gill, University of California at Riverside). To detect Na+,K+-ATPase, a mouse anti-chicken Na+,K+-ATPase -subunit monoclonal antibody, “a5”, developed by Dr. Douglas Fambrough was obtained from the Developmental Studies Hybridoma Bank under the auspices of the National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA. In Chapter 3, this antibody was used successfully to immunolabel Na+,K+-ATPase in the gills of the Atlantic stingray. Immunoblotting of V-H-ATPase B-subunit Immunoblots were performed on PVDF membranes that were prepared for Chapter 3, and contained 20 g of total gill membrane protein per lane. Details of the tissue preparation, electrophoresis, and blotting are described below. Preparation of tissue for immunoblots was similar to Claiborne et al. (1999), with modifications. On a single day, gill filaments from a freshwater, seawater-acclimated, and seawater stingray were prepared. First, the tissue was placed in ice cold

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72 homogenization buffer (250 mmol L-1 sucrose, 1 mmol L-1 Na-EDTA, 2 g mL-1 aprotinin, 2 g mL-1 leupeptin, 100 g mL-1 phenyl methylsulfonyl fluoride and 30 mmol L-1 Tris), and was homogenized with a Tissue-tearor (Biospec Products) in a 4C cold room on ice. Homogenates were filtered through cheese cloth and centrifuged (3000 g) for 5 min at 4C to remove nuclei and debris. The supernatant was then filtered through cheesecloth and centrifuged (50,000 g) for 30 min at 4C to pellet membrane fractions. The pellet was resuspended with a minimal volume of ice cold homogenization buffer, then an equal volume of a modified Laemmli sample buffer (Laemmli 1970), without bromophenol blue and -mercaptoethanol, was added to solubilize the proteins. The resulting protein samples were centrifuged for 5 to 10 s at 16,000 g to pellet any undissolved material. The total protein content of the supernatant was determined with a detergent compatible assay (Bio-Rad), after which bromophenol blue and -mercaptoethanol were added to final concentrations of 0.01% and 2%, respectively. A 20 g sample of total gill membrane protein from one stingray in each condition was loaded in triplicate and run on a 7.5% Tris-HCl precast polyacrylamide gel (Bio-Rad) for 1 h at 125 V. Proteins were then transferred to a PVDF membrane ( Bio-Rad) using a wet (20% MeOH, Tris-glycine) transfer unit for 2.5 h at 90 volts in a 4C cold room with stirring. The protein preparation, electrophoresis, and blotting were repeated for the remaining gill samples, which resulted in a total of five PVDF membranes with each containing, in triplicate, gill membrane protein from a single freshwater, seawater-acclimated, and seawater stingray. Because these PVDF membranes were previously immunostained (see Chapter 3), it was necessary to remove antibodies that were bound to the membrane with a strip

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73 buffer (62 mmol L-1 Tris-Base, 2% sodium lauryl sulfate, 0.6% -mercaptoethanol, pH 6.7). Each PVDF membrane was first soaked in 100% methanol for 15 minutes and then placed in the strip-buffer for 30 minutes at 60C. The PVDF was then placed in 3 washes of dH2O (5 minutes each) to remove any residual -mercaptoethanol. The PVDF was blocked with blocking buffer for 1.5 h at 25C, and then incubated with the rabbit anti-insect V-H-ATPase B-subunit polyclonal antibody (diluted 1:10,000 in blocking buffer) overnight at 4C. After antibody incubation, the PVDF was washed three times (15 min each) with TTBS, then incubated with an alkaline-phosphatase conjugated goat anti-rabbit IgG secondary antibody (Bio-Rad; diluted 1:3000 in blocking buffer) for 2 h at 25C. The PVDF was then washed three times (15 min each) with TTBS, and a substrate solution (Bio-Rad Immun-Star ECL Kit) was applied to the PVDF for 5 min at 25 C to initiate a luminescent signal. Binding of antibody was detected by exposing Hyperfilm-ECL imaging film (Amersham) to the PVDF membrane. Negatives were digitized into TIFF files using a flatbed scanner with transparency adapter (UMAX). As a control, stripped PVDF membranes were incubated with normal rabbit serum in blocking buffer, instead of the anti-V-H-ATPase polyclonal antibody. To quantify the relative abundance of V-H-ATPase immunoreactivity, I measured the optical density (uncalibrated) of the immunopositive band in each animal using Scion Image version 4.02 (Scion Corporation). On a given blot, optical density values were measured for each animal, and then standardized to the freshwater condition to calculate relative abundance. Therefore, all relative abundance measurements of freshwater gills

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74 were .0”, and those of seawater-acclimated and seawater gills were a fraction of the freshwater value. Immunohistochemical Localization of V-H-ATPase B-subunit The fixed gill tissue (stored in 75% ethanol) was dehydrated in an ethanol series, and embedded into paraffin wax. Serial sections of gill tissue, parallel to the long axis of the filament, were cut at 6 m and placed on poly-L-lysine coated slides (three sections per slide). Sections were deparaffinized in Hemo-De (Fisher Scientific), hydrated in an ethanol series, and washed in 10 mmol L-1 PBS. A hydrophobic PAP-Pen (Electron Microscopy Sciences) was used to draw circles around the tissue sections, and then 3% H2O2 was placed on the sections for 30 min to inhibit endogenous peroxidase activity. Sections were also blocked with Biogenex Protein Block (BPB; Normal goat serum with 1% bovine serum albumin, 0.09% NaN3, and 0.1% Tween-20) for 20 min. Then, the polyclonal rabbit anti-insect V-H-ATPase B-subunit antibody (diluted 1:10,000 in BPB) was incubated on the sections overnight at 4C. The antibody was rinsed off with PBS and then the sections were soaked in PBS for 5 min. The sections were incubated with a biotinylated goat anti-rabbit IgG secondary antibody (Biogenex) and a horseradish peroxidase-labeled strepavidin solution (Biogenex) for 20 min each at 25C (with a 5 min PBS wash after each incubation). Antibody binding was visualized by applying the chromagen 3,3’-diaminobenzidine tetra-hydrochloride (DAB, Biogenex) to the sections for 5 min at 25C. In each experiment, one section was exposed to normal rabbit serum or BPB in place of the anti-insect V-H-ATPase antibody as a negative control.

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75 Quantification of the immunohistochemistry was performed by counting the number of immunopositive (V-H-ATPase-rich) cells per gill lamella and per interlamellar space. For each animal, three immunostained slides were chosen randomly. On a section from each slide, the number of V-H-ATPase-rich cells was counted on 30 randomly selected lamellae and interlamellar regions. Lengths of lamellae were also measured to standardize lamellar cell counts. Results are expressed as number of V-H-ATPase-rich cells per 100 m of lamella, per interlamellar region, and per 100 m of lamella + interlamellar region (referred to as ‘sum’). Colocalization of V-H-ATPase with Na+,K+-ATPase To determine if V-H-ATPase and Na+,K+-ATPase were expressed in the same cells, I used a double-labeling technique modified from Verlander et al. (1996), consisting of sequential immunolocalization procedures using two different chromagens. Tissue sections for double-labeling were deparaffinized, hydrated, blocked, and stained for V-H-ATPase as described above. After treatment with a brown chromagen (DAB), the slides were rinsed in dH2O for 10 min and blocked with BPB for 20 min. A mouse anti-chicken Na+,K+-ATPase antibody (monoclonal antibody “a5” culture supernatant diluted 1:100 in BPB) was then applied to the sections overnight at 4C. Rinsing and developing were performed as described above, except a blue chromagen was used (Vector SG, Vector Laboratories, Burlingame, CA). Statistical Analyses Differences in relative abundances from immunoblots were detected using a Kruskal-Wallis non-parametric ANOVA, with a Kruskal-Wallis multiple comparisons test (Conover 1980). For mean number of V-H-ATPase-rich cells, differences among groups were detected using a one-way analysis of variance (ANOVA), with a Student

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76 Newman-Keuls multiple comparisons test. All tests were 2-tailed and differences were considered significant if P < 0.05. Results Immunoblotting of V-H-ATPase B-subunit The anti-insect V-H-ATPase B-subunit antibody bound to a 60.5 1.08 kDa protein in stingray gill membrane enrichments (Figure 4-1). No detectable signal was present when PVDF membranes were incubated with normal rabbit serum instead of the primary antibody (data not shown). Semi-quantitative immunoblotting showed that V-H-ATPase B-subunit immunoreactivity was highest in gills of freshwater stingrays, intermediate in seawater-acclimated, and lowest in seawater stingrays (Figure 4-2). Immunohistochemistry of V-H-ATPase B-subunit Regardless of salinity, immunohistochemical staining for the V-H-ATPase B-subunit occurred in relatively large cells of the gill epithelium, and did not appear to associate with pavement cells (Figures 4-3, 4-5). No staining was detected when BPB or normal rabbit serum was used instead of primary antibody (data not shown). The number of V-H-ATPase-rich cells in the gills was influenced by environmental salinity and followed a similar trend as V-H-ATPase immunoreactivity. The number of branchial V-H-ATPase-rich cells per 100 m of lamellae, per interlamellar region, and sum number of V-H-ATPase-rich cells (lamellar + interlamellar) was highest in freshwater stingrays, intermediate in seawater-acclimated, and lowest in seawater stingrays (Figure 4-4). Localization of the transporter within the V-H-ATPase-rich cells appeared to be qualitatively different among the three groups of stingrays. In freshwater stingrays, V-H-ATPase-rich cells were characterized by diffuse immunostaining throughout the cytoplasm and discrete basolateral localization (Figure 4-5 A). In V-H-ATPase-rich cells

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77 of seawater-acclimated and seawater stingrays, staining appeared to be darker in the cytoplasm with less discrete basolateral staining, relative to freshwater individuals (Figure 4-5). Discrete apical staining was not found in V-H-ATPase-rich cells from the gills of any stingray. Colocalization of V-H-ATPase Immunoreactivity with Na+,K+-ATPase Regardless of environmental salinity or location (lamellae vs. interlamellar region), expression of V-H-ATPase and Na+,K+-ATPase occurred in separate cells (Figure 4-6). It is interesting to note that V-H-ATPase-rich cells on gill lamellae of freshwater stingrays were usually located near the base of lamellae, while Na+,K+-ATPase-rich cells were primarily found on the distal parts of lamellae (Figure 4-6 A). The functional implications of this distribution are not known. On the interlamellar regions, V-H-ATPaseand Na+,K+-ATPase-rich cells were interspersed among each other (Figure 46). Discussion Results from this study are the first to show that V-H-ATPase expression in the gills of a euryhaline elasmobranch is influenced by environmental salinity. Immunoblotting with an anti-V-H-ATPase B-subunit antibody detected an approximate 60.5 kDa protein in the gill membrane enrichments (Figure 4-1), which is close to the expected size of 58 kDa reported for the ubiquitous B2 isoform (Nelson and Harvey 1999). Semi-quantitative immunoblotting showed that environmental salinity influenced the relative abundance of V-H-ATPase immunoreactivity in Atlantic stingray gills (Figure 4-2). In Chapter 3, I found a similar effect of environmental salinity on branchial Na+,K+-ATPase activity and immunoreactivity in Atlantic stingrays. These findings suggest that the gill epithelium of freshwater stingrays has an overall greater active transport potential

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78 compared to seawater-acclimated and seawater stingrays, possibly for the active uptake of NaCl and extrusion of H+/HCO3-. V-H-ATPase immunoreactivity in gills of seawater-acclimated stingrays was significantly lower than in gills of freshwater stingrays (Figure 4-2). This effect of salinity on branchial V-H-ATPase expression has also been reported in a teleost (O. mykiss), where gill V-H-ATPase activity and immunoreactivity decreased when freshwater trout were acclimated to seawater (Lin and Randall 1993, Lin et al. 1994). A lower branchial V-H-ATPase expression for the seawater-acclimated stingrays was expected, because active Na+ uptake would not be necessary in a seawater environment. Passive Na+/H+ exchangers are hypothesized to be responsible for acid extrusion in seawater fishes (Claiborne 1998, Claiborne et al. 1999, Edwards et al. 1999). The immunohistochemical results of this study suggest that the higher overall immunoreactivity of V-H-ATPase in freshwater stingray gills can be attributed to a greater number of V-H-ATPase-rich cells found in the gill epithelium, especially on the lamellae (Figures 4-3, 4-4). The dramatically lower number of V-H-ATPase-rich cells found on the lamellae of seawater-acclimated and seawater stingrays suggests that these cells may have a specialized function in freshwater stingrays, such as NaCl uptake. In Chapter 3, I reported a similar effect of environmental salinity on the number and distribution of Na+,K+-ATPase-rich cells in the gills of the Atlantic stingray. Localization of V-H-ATPase occurred in relatively large cells of the gill epithelium (Figures 4-3, 4-5) that are likely mitochondrion-rich, because V-H-ATPase-rich cells in transporting epithelia of other vertebrates are often mitochondrion-rich (Brown and Breton 1996), and V-H-ATPase-rich cells in the gills of a marine elasmobranch were

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79 mitochondrion-rich (Wilson et al. 1997). In freshwater stingrays, V-H-ATPase-rich cells were characterized by diffuse cytoplasmic and discrete basolateral immunostaining (Figure 4-5), which is opposite to the apical localization found in freshwater teleost gills (Lin et al. 1994, Sullivan et al. 1995, Wilson et al. 2000a). Basolateral localization of V-H-ATPase is relatively rare in vertebrates and to date has only been described in type B intercalated and mitochondrion-rich cells of the mammalian collecting duct and turtle urinary bladder (Stetson and Steinmetz 1985, Brown et al. 1988a, Brown et al. 1988b, Verlander et al. 1992, Verlander et al. 1994b, Brown and Breton 1996, Steinmetz et al. 1996). If the V-H-ATPase-rich cells of freshwater stingray gills are analogous in function to Type B intercalated cells then they would be involved with HCO3excretion and Cluptake via an apical Cl-/HCO3-exchanger (Weiner and Hamm 1990, Royaux et al. 2001, Tsuganezawa et al. 2001). This would be in contrast to freshwater teleost gills in which Cl-/HCO3exchange is thought to occur in Na+,K+-ATPase-rich chloride cells (Sullivan et al. 1996, Wilson et al. 2000a). In seawater-acclimated and seawater stingray gills, V-H-ATPase immunolabeling appeared to be stronger in the cytoplasm with less discrete basolateral localization, compared to freshwater individuals (Figure 4-5). Although ultrastructural studies would be required to verify these qualitative trends, the findings are consistent with V-H-ATPase regulation in other vertebrate tissues, in which recycling of the transporter between a cytoplasmic pool of vesicles and the plasma membrane was shown (Dixon et al. 1986, Stetson and Steinmetz 1986, Verlander et al. 1992, Verlander et al. 1994a, Brown and Breton 1996, Brown and Breton 2000). Therefore, the qualitative staining differences may indicate that seawater-acclimated and seawater stingrays have more V

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80 H-ATPase stored in cytoplasmic vesicles and less transporter on the basolateral membrane, compared to freshwater stingrays. This trend is consistent with the hypothesis that V-H-ATPase-rich cells secrete HCO3-, because Cl-/HCO3exchange in marine stingrays could be driven by the favorable gradient for Clto enter the cells from seawater, rather than the active generation of an outward HCO3gradient by basolateral V-H-ATPase. In seawater stingrays, V-H-ATPase-rich cells were only found on the interlamellar region of the gills, which corroborates the results of Wilson et al.(1997) who localized V-H-ATPase in the gills of a marine elasmobranch (S. acanthias) using an antibody to the A-subunit. Wilson et al. (1997) reported a cytoplasmic localization for V-H-ATPase in mitochondrion-rich cells, and suggested it was stored in vesicles that may be recruited to the apical membrane during a systemic acidosis (similar to a type Aintercalated cell). However, the results from this Chapter suggest that these vesicles would be recruited to the basolateral membrane during a systemic alkalosis (similar to a Type B intercalated cell). This would imply that another cell type and transporter is involved with acid excretion (see below). Because branchial V-H-ATPase-rich and Na+,K+-ATPase-rich cell (Chapter 3) abundance and distribution were affected similarly by salinity, I was interested in determining whether these two transporters localized to the same cells. Double-labeling gills for V-H-ATPase and Na+,K+-ATPase showed that these two transporters were expressed in separate cells, regardless of environmental salinity (Figure 4-6). This is important, because it suggests two types of ionocytes may exist in the gill epithelium of elasmobranch fishes. Previous studies have suggested the elasmobranch gill epithelium

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81 contained two types of mitochondrion-rich cells, based on the appearance of two distinct apical cell membrane morphologies (Laurent and Dunel 1980, Crespo 1982). This study provides the first immunohistochemical evidence for two mitochondrion-rich cell populations in the elasmobranch gill. The finding of separate V-H-ATPase-rich and Na+,K+-ATPase-rich cells has important functional implications. For example, this separation may indicate that Cluptake/HCO3excretion and Na+ uptake/H+ excretion occur in separate cells. Segregation of these processes is known to occur in the mammalian collecting duct and turtle urinary bladder. Type B intercalated cells express a basolateral V-H-ATPase that drives HCO3secretion and Cluptake via an apical Cl-/HCO3exchanger (Stetson et al. 1985, Stetson and Steinmetz 1985, Stetson and Steinmetz 1986, Brown et al. 1988a, Verlander et al. 1992). Type A intercalated cells express an apical V-H-ATPase that drives HCO3reabsorption via a basolateral band-3 (AE-1) Cl-/HCO3exchanger (Stetson and Steinmetz 1985, Stetson and Steinmetz 1986, Brown et al. 1988a, Verlander et al. 1988). Principal cells express basolateral Na+,K+-ATPase that drives Na+ uptake via an apical ENaC (Kashgarian et al. 1985, Wieczorek et al. 1999, Alvarez de la Rosa et al. 2000). I propose that the V-H-ATPase-rich cells in the stingray gill are functionally analogous to type B intercalated cells and are sites of Cluptake and HCO3excretion. I also propose that the Na+,K+-ATPase-rich cells are a functional amalgam of type A intercalated and principal cells, and are sites of H+ excretion and Na+ uptake. However, the Na+,K+-ATPase-rich cells probably use different apical mechanisms than the mammalian collecting duct and turtle urinary bladder. A likely candidate would be an apical NHE isoform that has been colocalized to Na+,K+-ATPase-rich cells in the gills of

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82 marine elasmobranchs (Edwards et al. 2002). A hypothetical model of NaCl and acid/base transport in the Atlantic stingray gill is presented in Figure 4-7. In summary, this study has shown that branchial V-H-ATPase expression in the Atlantic stingray is influenced by environmental salinity, with highest expression occurring in freshwater stingrays, followed by seawater-acclimated, and seawater individuals. The basolateral localization of V-H-ATPase suggests that it may be involved with active HCO3excretion and Cluptake, which is in contrast to the transporter’s proposed role of H+ excretion and Na+ uptake in freshwater teleost gills. The lack of apical V-H-ATPase immunostaining in the stingray gill suggests another cell type may be involved with acid excretion, possibly the Na+,K+-ATPase-rich cells. Results from this Chapter and Chapter 3 showed that a relatively high branchial expression of V-H-ATPase and Na+,K+-ATPase is associated with freshwater Atlantic stingrays, which suggests these two transporters might play an important role in the ability of this species to maintain NaCl and acid/base homeostasis in fresh water environments.

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83 kDa FW SWA SW 43 77 130 205 Figure 4-1. Representative immunoblot for V-H-ATPase B-subunit in gill membrane protein from freshwater (FW), seawater-acclimated (SWA), and seawater (SW) Atlantic stingrays. The antibody recognized a 60.5 1.08 kDa protein (arrow). Black lines represent migration of molecular weight markers (kDa).

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84 c b a SW SWA FW 1.2 1.0 0.8 0.6 0.4 0.2 0 Relative Abundance Figure 4-2. Relative abundance of immunoreactivity for 60.5 kDa protein representing the B-subunit of V-H-ATPase in the gill membrane enrichments of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) stingrays. Values are presented as means + 1 S.E. Lower case letters above error bars indicate statistical categorization of groups as determined by a Kruskal-Wallis multiple comparisons test (P < 0.05). Note that no error bars are present in FW, because of the standardization procedure (see Materials and Methods).

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85 C B A Figure 4-3. Representative photomicrographs of V-H-ATPase immunostaining in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). The V-H-ATPase-rich cells occurred on lamellae (finger-like projections) and/or interlamellar regions (basal to and between lamellae). Bar = 100 m.

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86 c b a A b c a C 12 10 8 6 4 2 0 b c a B S W F W SWA 5 4 3 2 1 0 6 Number of cells (per 100 m of lamella) 5 4 3 2 1 0 Number of cells (per interlamellar region) Number of cells (sum) Figure 4-4. Number of V-H-ATPase-rich cells in the gills of freshwater (FW), seawater-acclimated (SWA) and seawater (SW) Atlantic stingrays. Values are means + 1 S.E. Lower case letters above error bars indicate statistical categorization of the means as determined by a Student-Newman-Keuls multiple comparisons test (P < 0.05). Note that “Sum” = number of V-H-ATPase-rich cells per 100 m of lamella + per interlamellar region, and that both mean and S.E. for number of V-H-ATPase-rich cells per 100 m of lamella were .0” in SW stingrays.

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87 A B C Figure 4-5. Representative higher magnification photomicrographs of V-H-ATPase-rich cells in gills from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (1000x). Arrows indicate cells that best demonstrate qualitative differences in staining described in the text. Bar = 50 m.

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88 A B C Figure 4-6. Representative photomicrographs of V-H-ATPase immunolabeling (brown) colocalized with Na+,K+-ATPase (blue) in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). Note distinct brown and blue cells. Bar = 100 m.

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89 Fresh Water or Seawater Blood HCO3Na+ K+ H+ Cl? H+ Na+ ? Figure 4-7. Hypothetical model of NaCl and acid/base transport in the gills of the Atlantic stingray. Results from this study and Chapter 3 suggest that V-H-ATPase and Na+,K+-ATPase occur on basolateral regions of distinct mitochondrion-rich cell types. I hypothesize that the V-H-ATPase-rich cells act as HCO3secreting cells via an apical Cl-/HCO3exchanger that would also result in Cluptake. In contrast, I hypothesize that Na+,K+-ATPase-rich cells act as H+ secreting cells via an apical NHE that would also result in Na+ uptake. A “?” indicates that the transporter has not been detected in Atlantic stingrays.

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CHAPTER 5 IMMUNOCHEMICAL EVIDENCE OF A PENDRIN-LIKE CL-/HCO3EXCHANGER IN THE GILL EPITHELIUM OF THE ATLANTIC STINGRAY Introduction Pendrin is a recently discovered protein in mammals encoded by the Pendred disease syndrome (PDS) gene (Everett et al. 1997). It belongs to a diverse family of anion exchangers (SLC26) that includes such transporters as the sulfate-anion transporter (sat-1), diastrophic dysplasia sulfate transporter (DTDST), and congenital chloride diarrhea chloride-base exchanger (CLD) (Everett and Green 1999, Markovich 2001, Soleimani 2001). Pendrin was first described in the mammalian thyroid gland (Everett et al. 1997), and was discretely localized to the apical region of thyrocytes, where it is believed to function as a Cl-/iodide exchanger (Bidart et al. 2000, Royaux et al. 2000, Lacroix et al. 2001). This transporter has Cl-/iodide, Cl-/HCO3-, Cl-/OH-, and Cl-/formate exchange activity when expressed in Xenopus oocytes and HEK-293 cells (Scott et al. 1999, Scott and Karniski 2000, Soleimani et al. 2001). Recent studies have detected both pendrin mRNA (Royaux et al. 2001, Soleimani et al. 2001) and protein (Royaux et al. 2001) in the cortical collecting duct (CCD) of the mammalian nephron. Royaux et al. (2001) localized pendrin to the apical region of vacuolar-proton-ATPase (V-H-ATPase)-rich intercalated cells that did not have basolateral anion-exchanger (AE)-1 immunoreactivity. Furthermore, Royaux et al. (2001) showed that isolated, perfused CCD tubules from HCO3loaded pendrin-knockout mice had a decreased capacity for HCO3secretion compared to CCD tubules from wild90

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91 type mice. Taken together, these data not only suggested that intercalated cells with pendrin were HCO3secreting cells, but also that pendrin may represent the apical Cl-/HCO3exchanger in type-B intercalated cells. In elasmobranchs, the gills are the primary site of acid/base-related ion transport (Heisler 1988). However, no study has determined the identity or cellular location of a Cl-/HCO3exchanger in the elasmobranch gill, although net HCO3excretion is known to occur across this epithelium (Holder et al. 1955, Swenson and Maren 1987). In the gills of a euryhaline elasmobranch (Atlantic stingray, Dasyatis sabina), I have previously identified cells that are rich in basolateral V-H-ATPase, which may be functionally analogous to HCO3secreting cells of the mammalian CCD (Chapter 4). Therefore, I hypothesized that cells with basolateral V-H-ATPase in the stingray gill may have apical pendrin to mediate Cl-/HCO3exchange. To date, no studies of pendrin in the elasmobranch gill, or in transport epithelia from any lower vertebrate, have been reported. The Atlantic stingray is unique because it is one of the few elasmobranch species with the physiological capability of living in both fresh and seawater environments. In Chapters 3 and 4, I showed that both Na+,K+-ATPase and V-H-ATPase are expressed at relatively high levels in gills of freshwater Atlantic stingrays, and subsequently decrease when the animals acclimate to seawater. To my knowledge, the influence of environmental salinity on expression of a branchial Cl-/HCO3exchanger has not been examined in any fish. Understanding how salinity influences expression of an anion exchanger is especially of interest in the Atlantic stingray, because it may reveal mechanisms this species uses for NaCl and acid/base regulation in fresh and seawater

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92 environments. For example, evidence in freshwater teleosts suggests that an AE-1-like Cl-/HCO3exchanger (Sullivan et al. 1996, Wilson et al. 2000a) contributes to HCO3secretion for acid/base balance and to Cluptake for NaCl homeostasis (Perry et al. 1981, Perry and Randall 1981, Laurent and Perry 1990, Perry et al. 1992, Goss and Perry 1994, Perry and Goss 1994). In seawater teleost gills, Cl-/HCO3exchanger activity persists for acid/base regulation (Claiborne et al. 1997), but could potentially complicate NaCl balance by leading to Claccumulation (Claiborne 1998). The goals of this study were to determine if pendrin is expressed in the gills of the Atlantic stingray, to identify the specific cellular location of pendrin in the gill epithelium, and to determine if expression of pendrin in the gills is influenced by environmental salinity. My results present the first evidence of pendrin expression in a lower vertebrate, and suggest that pendrin or a pendrin-like Cl-/HCO3exchanger exists on the apical membrane of V-H-ATPase-rich cells in the Atlantic stingray gill epithelium. Furthermore, branchial pendrin expression is decreased by increased environmental salinity. Materials and Methods Animal Collection and Holding Conditions Ten Atlantic stingrays were captured from the St. Johns River, FL (Lake Jesup or Lake George) with trot lines, transported to Gainesville, FL, and held in two 379 L freshwater, closed-system tanks (five rays per tank; < 1 ppt salinity). In addition, five marine Atlantic stingrays were captured via hook and line from Cedar Key, FL, transported to Gainesville, FL, and held in a 379 L seawater, closed-system tank (32 ppt). Five of the freshwater stingrays were left in fresh water (referred to as freshwater stingrays), while the other five were gradually acclimated to seawater as follows (referred

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93 to as seawater-acclimated stingrays). After one week in fresh water, the salinity was raised to 16 ppt over 2 days (8 ppt per day). After two days in 16 ppt, the salinity was raised to 32 ppt seawater over three days. The animals remained in 32 ppt seawater for one week before tissue samples were taken. The marine stingrays from Cedar Key remained in 32 ppt seawater for the entire period (referred to as seawater stingrays). All animals were fed live grass shrimp (Palaemonetes sp.) every other day, and were starved 48 h before tissue collection. Water temperature of all tanks was maintained at 25C, and pH was adjusted to 8.2 using freshwater and marine buffers (Seachem). The tanks were also equipped with biological filtration, which maintained ambient NH3 and NO3 levels below 1 part per million. Collection of Gill Tissue Animals were anesthetized in 4 L of a 0.01% 3-aminobenzoic acid ethyl ester (MS-222, Sigma) solution made with tank water. For freshwater stingrays, the solution was buffered with a commercial freshwater pH buffer (Seachem) to prevent acidification by the MS-222. Once anesthetized, animals were placed ventral side up in a slanted water bath with their gills immersed in the anesthetic. In order to clear the gills and rectal gland of red blood cells, the animals were perfused with a marine elasmobranch Ringer's solution at 4C (Forster et al. 1972). However, for freshwater stingrays, NaCl, urea, and trimethylamine oxide concentrations in the Ringer’s were reduced to 200, 200, and 41 mmol L-1, respectively. The skin ventral to the heart and pericardium was removed, and 0.5 to 1.0 ml of blood was removed from the ventricle with a heparinized 25 gauge needle attached to a 1 ml syringe. An equal volume of heparinized Ringer's solution was then injected into the

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94 ventricle and allowed to circulate for a few minutes. A cannula, connected to a perfusion bottle (1 m above animal), was inserted into the conus arteriosus and held by forceps. Once the perfusion was started, the sinus venosus was cut to relieve backpressure. The perfusion was continued until the gills appeared bleached and the fluid exiting the pericardial cavity was clear of blood (usually 3 to 5 min). Immediately after the perfusion, the animal was pithed, and the second left and right gill arches were removed and placed in an elasmobranch Ringer's solution on ice. For immunohistochemistry, gill filaments were trimmed off the arches and placed in fixative (3% paraformaldehyde, 0.05% glutaraldehyde, 0.05% picric acid in 10 mmol L-1 phosphate buffered saline, pH 7.3) for 24 h at 4C, then transferred to two changes of 75% ethanol for removal of fixative. Tissues were left in the second change of 75% ethanol until embedded. Additional filaments were snap frozen in liquid nitrogen for immunoblot analysis and stored at -80C until analyzed. Antibodies The antibody used to detect pendrin was developed by Royaux et al. (2000) and was kindly provided by Dr. Ines Royaux, National Institutes of Health. This antibody is an affinity purified, rabbit polyclonal antibody raised against amino acids 630 to 643 of human pendrin. This sequence of amino acids is near the carboxy terminus of pendrin and is completely conserved among humans, rats, and mice (Everett et al. 1999). The antibody has been used to immunolabel pendrin in HEK293 cells transfected with pendrin cDNA and in rat thyroid tissue (Royaux et al. 2000). To detect Na+,K+-ATPase, a mouse anti-chicken Na+,K+-ATPase -subunit monoclonal antibody, “a5”, developed by Dr. Douglas Fambrough was obtained from the Developmental Studies Hybridoma

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95 Bank under the auspices of the National Institute of Child Health and Human Development and maintained by The University of Iowa, Department of Biological Sciences, Iowa City, IA 52242. To detect V-H-ATPase, a rabbit anti-insect V-H-ATPase B-subunit polyclonal antibody was kindly provided by Dr. William Harvey, Whitney Laboratory, University of Florida, with permission from Dr. Sarjeet Gill, University of California at Riverside. In Chapters 3 and 4, I have successfully used the latter two antibodies to detect Na+,K+-ATPase and V-H-ATPase in gills of the Atlantic stingray. Immunoblot Analysis of Pendrin Immunoreactivity Immunoblots were performed on PVDF membranes that were prepared for Chapter 3, and contained 20 g of total gill membrane protein per lane. Details of the tissue preparation, electrophoresis, and blotting are described below. Preparation of tissue for immunoblots was similar to Claiborne et al. (1999), with modifications. On a single day, gill filaments from a freshwater, seawater-acclimated, and seawater stingray were prepared. First, the tissue was placed in ice cold homogenization buffer (250 mmol L-1 sucrose, 1 mmol L-1 Na-EDTA, 2 g mL-1 aprotinin, 2 g mL-1 leupeptin, 100 g mL-1 phenyl methylsulfonyl fluoride and 30 mmol L-1 Tris), and was homogenized with a Tissue-tearor (Biospec Products) in a 4C cold room on ice. Homogenates were filtered through cheese cloth and centrifuged (3000 g) for 5 min at 4C to remove nuclei and debris. The supernatant was then filtered through cheesecloth and centrifuged (50,000 g) for 30 min at 4C to pellet membrane fractions. The pellet was resuspended with a minimal volume of ice cold homogenization buffer, then an equal volume of a modified Laemmli sample buffer (Laemmli 1970), without bromophenol blue and -mercaptoethanol, was added to solubilize the proteins.

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96 The resulting protein samples were centrifuged for 5 to 10 s at 16,000 g to pellet any undissolved material. The total protein content of the supernatant was determined with a detergent compatible assay (Bio-Rad), after which bromophenol blue and -mercaptoethanol were added to final concentrations of 0.01% and 2%, respectively. A 20 g sample of total gill membrane protein from one stingray in each condition was loaded in triplicate and run on a 7.5% Tris-HCl precast polyacrylamide gel (Bio-Rad) for 1 h at 125 V. Proteins were then transferred to a PVDF membrane ( Bio-Rad) using a wet (20% MeOH, Tris-glycine) transfer unit for 2.5 h at 90 volts in a 4C cold room with stirring. The protein preparation, electrophoresis, and blotting were repeated for the remaining gill samples, which resulted in a total of five PVDF membranes with each containing, in triplicate, gill membrane protein from a single freshwater, seawater-acclimated, and seawater stingray. Because these PVDF membranes were previously immunostained (see Chapters 3 and 4), it was necessary to remove antibodies that were bound to the membrane with a strip-buffer (62 mmol L-1 Tris-Base, 2% sodium lauryl sulfate, 0.6% -mercaptoethanol, pH 6.7). Each PVDF membrane was first soaked in 100% methanol for 15 minutes and then placed in the strip-buffer for 30 minutes at 60C. The PVDF was then placed in 3 washes of dH2O (5 minutes each) to remove any residual -mercaptoethanol. The PVDF was blocked with blocking buffer for 1.5 h at 25C, and then incubated with the rabbit anti-human pendrin polyclonal antibody (diluted 1:5000 in blocking buffer) overnight at 4C. After antibody incubation, the PVDF was washed three times (15 min each) with TTBS, then incubated with an alkaline-phosphatase conjugated goat anti-rabbit IgG

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97 secondary antibody (Bio-Rad; diluted 1:3000 in blocking buffer) for 2 h at 25C. The PVDF was then washed three times (15 min each) with TTBS, and a substrate solution (Bio-Rad Immun-Star ECL Kit) was applied to the PVDF for 5 min at 25 C to initiate a luminescent signal. Binding of antibody was detected by exposing Hyperfilm-ECL imaging film (Amersham) to the PVDF membrane. Negatives were digitized into TIFF files using a flatbed scanner with transparency adapter (UMAX). As a control, stripped PVDF membranes were incubated with normal rabbit serum in blocking buffer, instead of the anti-pendrin polyclonal antibody. To quantify the relative abundance of pendrin immunoreactivity, I measured the optical density (uncalibrated) of the immunopositive band in each animal using Scion Image version 4.02 (Scion Corporation). On a given blot, optical density values were measured for each animal, and then standardized to the freshwater condition to calculate relative abundance. Therefore, all relative abundance measurements of freshwater gills were .0”, and those of seawater-acclimated and seawater gills were a fraction of the freshwater value. Immunohistochemical Localization of Pendrin Immunoreactivity The fixed gill tissue (stored in 75% ethanol) was dehydrated in an ethanol series, and embedded into paraffin wax. Serial sections of gill tissue, parallel to the long axis of the filament, were cut at 6 m and placed on poly-L-lysine coated slides (three sections per slide). Sections were deparaffinized in Hemo-De (Fisher Scientific), hydrated in an ethanol series, and washed in 10 mmol L-1 PBS. A hydrophobic PAP-Pen (Electron Microscopy Sciences) was used to draw circles around the tissue sections, and then 3% H2O2 was placed on the sections for 30 min to inhibit endogenous peroxidase activity.

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98 Sections were also blocked with Biogenex Protein Block (BPB; Normal goat serum with 1% bovine serum albumin, 0.09% NaN3, and 0.1% Tween-20) for 20 min. Then, the polyclonal rabbit anti-human pendrin antibody (diluted 1:1500 in BPB) was incubated on the sections overnight at 4C. The antibody was rinsed off with PBS and then the sections were soaked in PBS for 5 min. The sections were incubated with a biotinylated goat anti-rabbit IgG secondary antibody (Biogenex) and a horseradish peroxidase-labeled strepavidin solution (Biogenex) for 20 min each at 25C (with a 5 min PBS wash after each incubation). Antibody binding was visualized by applying the chromagen 3,3’-diaminobenzidine tetra-hydrochloride (DAB; Biogenex) to the sections for 5 min at 25C. In each experiment, one section was exposed to normal rabbit serum or BPB in place of the anti-pendrin antibody as a negative control. Colocalization of Pendrin Immunoreactivity with V-H-ATPase and Na+,K+-ATPase To determine if pendrin immunoreactivity was expressed in V-H-ATPase-rich and/or Na+,K+-ATPase-rich cells, I used a double-labeling technique consisting of sequential immunolocalization procedures using two different chromagens. Tissue sections for double-labeling were first deparaffinized, hydrated, blocked, and stained for pendrin as described above (except a 1:1000 dilution of the anti-pendrin antibody was used when colocalizing with V-H-ATPase). After treatment with the brown chromagen (DAB), the slides were rinsed in dH2O for 10 min and blocked with BPB for 20 minutes. Then, an antibody to V-H-ATPase (polyclonal antibody serum diluted 1:5000 in BPB) or Na+,K+-ATPase (monoclonal antibody “a5” culture supernatant diluted 1:100 in BPB) was applied to the sections overnight at 4C. Rinsing and developing was performed as described above, except a blue chromagen was used (Vector SG, Vector Laboratories).

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99 Statistical Analyses Differences in pendrin immunoreactivity relative abundance measurements were detected using a Kruskal-Wallis non-parametric ANOVA, with a Kruskal-Wallis multiple comparisons test (Conover 1980). All tests were 2-tailed and differences were considered significant at P < 0.05. Results Immunoblot Analysis of Pendrin Immunoreactivity In immunoblots of membrane proteins isolated from stingray gills, under all salinities tested, the anti-human pendrin antibody bound to a protein of 143.7 2.8 kDa (Figure 5-1). No detectable signal was present when PVDF membranes were incubated with normal rabbit serum instead of the primary antibody (data not shown). Semi-quantitative immunoblotting revealed that the relative abundance of pendrin immunoreactivity was highest in gill membrane protein from freshwater stingray gills (Figure 5-2). In gill membrane protein from seawater-acclimated stingrays, pendrin immunoreactivity was diminished compared to freshwater gill protein, and was comparable to that from seawater stingrays (Figure 5-2). Immunohistochemical Localization of Pendrin Immunoreactivity In freshwater stingrays, numerous pendrin-positive cells were found on both gill lamellae and interlamellar regions (Figure 5-3 A). In seawater-acclimated stingray gills, pendrin-positive cells were primarily detected on interlamellar regions, and were infrequently found on lamellae (Figure 5-3 B). In seawater stingrays, pendrin-positive cells were found exclusively on interlamellar regions, with no detectable immunoreactivity on lamellae (Figure 5-3 C). No staining was detected when non-immune rabbit serum or BPB was used instead of primary antibody (data not shown).

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100 Higher magnification images of pendrin-positive cells revealed that pendrin immunoreactivity was localized to the apical region (Figure 5-4), regardless of salinity. However, qualitative differences in immunostaining existed among the three stingray groups. In freshwater stingray gills, localization of pendrin immunoreactivity was most apical, discrete, and intense (Figure 5-4 A). In contrast, pendrin immunostaining in seawater-acclimated and seawater stingray gills was diffuse throughout the subapical cytoplasm, and did not have discrete apical pendrin immunoreactivity (Figure 5-4 B, C). Colocalization of Pendrin Immunoreactivity with V-H-ATPase and Na+,K+-ATPase Double-labeling of gills for pendrin immunoreactivity and V-H-ATPase revealed a similar distribution for these two transporters (Figure 5-5). Regardless of environmental salinity, all pendrin immunolabeling occurred in the apical region of cells that stained for V-H-ATPase (Figure 5-6). Double-labeling of pendrin immunoreactivity and Na+,K+-ATPase showed that these two transporters did not have a similar distribution (Figure 5-7) and occurred in separate cells (Figure 5-8), regardless of environmental salinity. Discussion The findings from this study present the first evidence of a pendrin-like transporter in an ion transporting tissue from any lower vertebrate. Immunoblotting with an anti-human pendrin antibody detected the presence of an approximate 144 kDa protein in gill membrane enrichments from Atlantic stingrays (Figure 5-1). This is slightly greater than the reported size of 100 kDa for pendrin in mammals (Royaux et al. 2000, Soleimani et al. 2001). The difference in size may indicate that the pendrin-like protein in elasmobranchs is composed of more amino acids, and/or that the protein is heavily glycosylated in elasmobranchs, compared to mammals. Pendrin has two potential glycosylation sites in mammals (Everett et al. 1997), but the protein has not been

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101 sequenced in elasmobranchs, therefore the number of amino acids and potential glycosylation sites in the elasmobranch protein are unknown. Semi-quantitative immunoblotting revealed that pendrin immunoreactivity was most abundant in gill tissue from freshwater stingrays, compared to seawater-acclimated and seawater stingrays (Figure 5-2). Therefore, the freshwater stingray gill epithelium appears to have a relatively high potential for pendrin-like Cl-/HCO3exchange. Greater expression of a Cl-/HCO3exchanger would be physiologically favorable in a freshwater environment, because increased Cluptake would be necessary to counteract diffusional and urinary losses of Clto the environment. Uptake of Clvia a pendrin-like transporter would also provide a route for HCO3secretion that would contribute to acid/base regulation. The immunohistochemical findings showed that the distribution of cells with detectable pendrin immunoreactivity was influenced by environmental salinity. In freshwater stingray gills, pendrin-positive cells were found on both gill lamellae and interlamellar regions, whereas pendrin-positive cells were primarily found in the interlamellar regions of seawater-acclimated and seawater stingray gills (Figure 5-3). Furthermore, the intensity of the immunolabeling appeared to be stronger in pendrin-positive cells of freshwater stingray gills, relative to seawater-acclimated and seawater stingrays (Figure 5-4). These differences in cellular distribution and intensity are consistent with the greater pendrin immunoreactivity detected in freshwater stingray gills by immunoblotting, compared to gills of seawater-acclimated and seawater stingrays (Figure 5-2).

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102 Pendrin immunohistochemistry also showed that localization of pendrin immunoreactivity within pendrin-positive cells occurred in the apical region, and that this subcellular localization was influenced by environmental salinity. In freshwater stingray gills, pendrin-positive cells had discrete, intense apical localization (Figure 5-4 A). In seawater-acclimated and seawater stingray gills, pendrin-positive cells lacked discrete apical staining, and instead exhibited weaker, diffuse staining throughout the subapical cytoplasm (Figure 5-4 B, C). These observations are similar to findings reported for transporters in other ion-secreting epithelia that are trafficked between a cytoplasmic pool of vesicles and the plasma membrane (Dixon et al. 1986, Verlander et al. 1992, Verlander et al. 1994a, Lehrich et al. 1998). Therefore, the differences that I observed may indicate that freshwater stingray gills have more pendrin-like transporters inserted into the apical membrane of pendrin-positive cells than seawater-acclimated and seawater stingrays to mediate apical Cl-/HCO3-exchange. This would be consistent with the physiologic need for enhanced Cluptake in freshwater environments to maintain NaCl balance. In seawater-acclimated and seawater stingrays, less apical insertion of a pendrin-like transporter may be adequate, because Cl-/HCO3-exchange would only be required for HCO3secretion to maintain acid/base balance; uptake of Clis no longer physiologically necessary in seawater environments and would actually be opposite to the needs of NaCl homeostasis. To date, pendrin-trafficking has not been described in mammalian tissues; this is the first suggestion of such regulation for this transporter. Double-labeling experiments clearly showed that pendrin expression was exclusively found in the apical region of V-H-ATPase-rich cells and not Na+,K+-ATPase-rich cells (Figures 5-5 to 5-8). These results are similar to pendrin localization reported

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103 in the mammalian CCD, where pendrin immunolabeling occurred in the apical region of HCO3secreting V-H-ATPase-rich intercalated cells that were not immunoreactive for AE-1 (Royaux et al. 2001). Because V-H-ATPase-rich cells in the Atlantic stingray gill have basolateral localization of V-H-ATPase (Chapter 4) and have apical pendrin immunolabeling (Figure 5-6), I hypothesize that these cells are analogous in function to type B intercalated cells of the mammalian CCD and are a site of apical Cl-/HCO3exchange, as proposed in Chapter 4. In particular, I propose that a basolateral V-H-ATPase could establish a favorable electrochemical gradient for HCO3secretion that would drive apical Cl-/HCO3exchange via a pendrin-like transporter. This is especially important in freshwater stingrays, because ambient NaCl concentration is low and passive uptake of Clcould not drive apical Cl-/HCO3exchange activity. However, in seawater stingrays, ambient NaCl concentrations are high and passive uptake of Clcould drive Cl-/HCO3exchange activity with less dependence upon a basolateral V-H-ATPase. Therefore, the salinity related differences in branchial expression of both pendrin immunoreactivity and V-H-ATPase (see Chapter 4) in the Atlantic stingray are consistent with the model of NaCl and acid/base transport mechanisms in fresh and seawater elasmobranch gills that I originally proposed in Chapter 4. I have incorporated my current results and those from a recent study on Na+/H+ exchangers in elasmobranch gills (Edwards et al. 2002) to further develop my model of ion transport in the elasmobranch gill epithelium (Figure 5-9). In summary, I have shown that pendrin immunoreactivity is present in the gills of the Atlantic stingray, which is the first evidence of a pendrin-like transporter in any tissue from a lower vertebrate. Pendrin immunoreactivity is most abundant and most apical in

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104 the gills of freshwater stingrays, compared to seawater-acclimated and seawater stingrays, and only occurs in the apical region of V-H-ATPase-rich cells, regardless of salinity. In conclusion, my findings suggest that a pendrin-like transporter may contribute to apical Cl-/HCO3exchange in the gill epithelium of the Atlantic stingray and may play an important role in the mechanisms of NaCl and acid/base homeostasis that allow this euryhaline species to inhabit both fresh and seawater environments.

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105 kDa SW SW SWA SWA FW FW 51 91 119 206 Figure 5-1. Representative immunoblot for pendrin immunoreactivity in gill membrane protein from freshwater (FW), seawater-acclimated (SWA), and seawater (SW) Atlantic stingrays. Black lines represent migration of molecular weight markers (kDa). The anti-human pendrin antibody recognized an approximate 144 kDa protein (arrow).

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106 1.2 1.2 b b b b a a 1.0 1.0 Relative Abundance 0.8 0.8 0.6 0.6 0.4 0.4 0.2 0.2 0 0 FW FW SWA SWA SW SW Figure 5-2. Relative abundance of immunoreactivity for 144 kDa band representing pendrin, in gill membrane protein of freshwater (FW), seawater-acclimated (SWA), and seawater (SW) Atlantic stingrays. N = 5 for all groups. Values are presented as means + 1 S.E. Lower case letters above error bars indicate statistical categorization of groups as determined by a Kruskal-Wallis multiple comparisons test (P < 0.05). Note that no error bars are present in FW, because of the standardization procedure (see Materials and Methods).

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107 A A B B C C Figure 5-3. Representative photomicrographs of pendrin immunolabeling in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). Pendrin-positive cells (brown) occurred on lamellae (finger-like projections) and/or interlamellar regions (basal to and between lamellae). Scale bar = 100 m.

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108 A B C Figure 5-4. Higher magnification photomicrographs of pendrin immunolabeling in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (1000x). Arrows indicate apical regions of cells that best demonstrate the qualitative differences described in text. Scale bar = 50 m.

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109 A B C Figure 5-5. Representative photomicrographs of pendrin immunolabeling (brown) colocalized with V-H-ATPase (blue) in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). Scale bar = 100 m.

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110 A B C Figure 5-6. Higher magnification photomicrographs of pendrin immunolabeling (brown) colocalized with V-H-ATPase (blue) in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (1000x). Scale bar = 50 m.

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111 B A C Figure 5-7. Representative photomicrographs of colocalization of pendrin immunolabeling (brown) colocalized with Na+,K+-ATPase (blue) in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (400x). Scale bar = 100 m.

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112 A B C Figure 5-8. Higher magnification photomicrographs of pendrin immunolabeling (brown) colocalized with Na+,K+-ATPase (blue) in longitudinal sections of gill filaments from freshwater (A), seawater-acclimated (B), and seawater (C) Atlantic stingrays (1000x). Thin arrows indicate apical region of pendrin-positive cells, and wide arrows indicate basolateral region of Na+,K+-ATPase-rich cells. Scale bar = 50 m.

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113 FreshWater Blood H+ H+H+ Na+?HCO3-ClPDSHCO3-ClPDSHCO3-ClPDSNa+K+ Na+K+ H+ H+ H+ H+ SeawaterBlood HCO3-ClPDSNa+K+ Na+K+ H+ H+H+ Na+NHE Figure 5-9. Model of NaCl and acid/base regulation in the freshwater (left) and seawater (right) elasmobranch gill. In freshwater elasmobranchs, I propose that V-H-ATPase-rich cells are sites of Cluptake and HCO3secretion via an apical pendrin-like Cl-/HCO3exchanger. The basolateral V-H-ATPase would actively pump H+ across the basolateral membrane to create a favorable electrochemical gradient for HCO3secretion and Cluptake via a pendrin-like transporter. I hypothesize that Na+,K+-ATPase-rich cells are a site of Na+/H+ exchange in freshwater elasmobranchs, but an apical transporter responsible for such an exchange has not been identified to date. In seawater elasmobranchs, I propose that V-H-ATPase-rich cells function similarly to those in gills of freshwater elasmobranchs. However, less plasma membrane insertion of apical pendrin-like Cl-/HCO3exchangers and basolateral V-H-ATPase would be adequate, because no physiological need exists for Cluptake, and Cl-/HCO3exchange can occur more efficiently in a high NaCl environment, respectively. I hypothesize that the Na+,K+-ATPase-rich cells are a site of Na+/H+ exchange via apical Na+/H+ exchangers that were localized to Na+,K+-ATPase-rich cells by Edwards et al. (2002). “NHE” = Na+/H+ exchanger, “PDS” = pendrin-like Cl-/HCO3exchanger. A “?” indicates that the transporter has not been detected. Note that depicted differences in plasma membrane insertion of PDS and V-H-ATPase between fresh and seawater are based on qualitative immunohistochemical observations from this study and Chapter 4.

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CHAPTER 6 SUMMARY AND FUTURE DIRECTIONS Summary Relatively few studies exist on the mechanisms of osmotic and acid/base balance in elasmobranch fishes, compared to the abundance of literature on fresh and seawater teleost fishes. Before this dissertation, studies on elasmobranch osmoregulation were usually limited to measurements of plasma osmotic parameters and rectal gland function in stenohaline marine species, because euryhaline or freshwater elasmobranch species are uncommon. The goals of this dissertation were to describe the general osmoregulatory parameters of a euryhaline elasmobranch and determine some of the extra-renal mechanisms it used to maintain NaCl and acid/base balance in marine and freshwater environments. In Chapter 2, I measured plasma osmolytes and rectal gland weights of freshwater Atlantic stingrays (Dasyatis sabina)living in the St. Johns River, FL, and determined how these parameters changed after acclimation to seawater. I hypothesized that the freshwater stingrays may show physiological divergence from marine Atlantic stingrays, because the freshwater individuals reproduce and complete their life cycle in the river. I found that freshwater Atlantic stingrays were characterized by reduced plasma concentrations of major osmolytes (e.g., NaCl and urea) and a smaller rectal gland, compared to marine Atlantic stingrays. However, the freshwater stingrays still maintained a substantial hyperosmotic gradient (approximately 600 mOsm kg-1) between their plasma and the freshwater environment. This osmoregulatory strategy is similar to 114

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115 other euryhaline elasmobranchs in freshwater, such as the bull shark (Carcharhinus leucas). When freshwater Atlantic stingrays were acclimated to seawater (seawater-acclimated stingrays), their plasma osmotic parameters were similar to typical marine stingrays, despite no change in rectal gland size. These results suggested that the freshwater populations were not physiologically restricted to the St. Johns River and could potentially migrate to seawater. In Chapter 3, I addressed the cellular mechanisms involved with NaCl regulation in the Atlantic stingray by examining how environmental salinity influenced Na+,K+-ATPase expression in the gills and rectal gland. The Na+,K+-ATPase plays a key role in energizing active NaCl excretion in the marine elasmobranch rectal gland and gills of marine teleosts, and may be important for active NaCl uptake in gills of freshwater teleosts. Using a ouabain-specific ATPase assay and immunoblotting I found that gills of freshwater stingrays had the highest activity and immunoreactivity of Na+,K+-ATPase, compared to gills of seawater-acclimated and seawater stingrays. Using immunohistochemistry, I found that freshwater stingray gills also had the greatest number of Na+,K+-ATPase-rich cells, compared to seawater-acclimated and seawater stingrays. In rectal glands, Na+,K+-ATPase activity and immunoreactivity were lowest in freshwater stingrays, compared to seawater-acclimated and seawater individuals. The results from this study suggested that the gills played an important role in active transport for freshwater stingrays, possibly for NaCl uptake and acid/base regulation. In contrast, the results suggested that rectal glands played a more important role in seawater-acclimated and seawater stingrays, possibly for active NaCl excretion. This study was first to show

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116 that environmental salinity influenced activity and expression of Na+,K+-ATPase in the gills and rectal gland of an elasmobranch. In Chapter 4, I continued my investigation of the cellular mechanisms involved with extra-renal ion regulation in the Atlantic stingray, but focused on the gills. I examined how salinity influenced the expression of vacuolar proton-ATPase (V-H-ATPase), which is a transporter known to have a function in both NaCl and acid/base homeostasis. In the gills of freshwater teleost fishes, V-H-ATPase is found on the apical membrane of pavement and Na+,K+-ATPase-rich cells, where it energizes H+ excretion and Na+ uptake. In marine elasmobranch gills, this transporter was identified in mitochondrion-rich cells, but its expression in freshwater individuals had not been examined. Using immunoblotting and immunohistochemistry, I found that gills from freshwater stingrays had the greatest immunoreactivity of V-H-ATPase and greatest number of V-H-ATPase-rich cells, compared to seawater-acclimated and seawater stingrays. The V-H-ATPase-rich cells were characterized by basolateral immunolabeling that was more basolateral and discrete in freshwater stingray gills, compared to seawater-acclimated and seawater stingrays. Double-label immunohistochemistry showed that V-H-ATPase-rich cells were distinct from Na+,K+-ATPase-rich cells, which suggested two types of mitochondrion-rich cells existed in the elasmobranch gill epithelium. Based on these findings, I proposed a model of NaCl and acid/base regulation in the Atlantic stingray gill where V-H-ATPase-rich cells and Na+,K+-ATPase-rich cells were sites of Cluptake/HCO3excretion and Na+ uptake/H+ excretion, respectively. The results from this chapter were the first to show that environmental salinity influenced expression of V-H-ATPase in the gills of an elasmobranch.

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117 In Chapter 5, I completed my study on the branchial mechanisms of NaCl and acid/base regulation in the Atlantic stingray. I examined how salinity influenced expression of pendrin, which is a Cl-/HCO3exchanger that has recently been described in the mammalian kidney. Before this study, pendrin had not been investigated in ion regulatory tissues of any lower vertebrate. Using immunoblotting and immunohistochemistry, I found that pendrin immunoreactivity was present in the gills of the Atlantic stingray and was influenced by environmental salinity. Pendrin immunoreactivity and pendrin-positive cells were most abundant in gills of freshwater stingrays, compared to seawater acclimated and seawater individuals. Pendrin-positive cells were characterized by apical immunolabeling, and the localization was more apical, discrete, and intense in freshwater stingray gills, compared to seawater-acclimated and seawater individuals. Double-label immunohistochemistry showed that pendrin immunoreactivity was localized to the apical region of V-H-ATPase-rich cells, but not Na+,K+-ATPase-rich cells. This suggested that V-H-ATPase-rich cells were a site of Cl-/HCO3exchange mediated by an apical pendrin-like transporter, which is consistent with the original model of NaCl and acid/base regulation that I proposed in Chapter 4. Results from this chapter were the first to show pendrin immunoreactivity in a lower vertebrate, and the first to show that salinity influences expression of a Cl-/HCO3exchanger in the gills of any fish. Future Directions I feel that the major contributions of this dissertation are that I have established an excellent model species to study extra-renal mechanisms of elasmobranch ion regulation (the Atlantic stingray, D. sabina), and I have presented a model of ion transport in the Atlantic stingray gill that can explain how this species maintains NaCl and acid/base

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118 homeostasis in marine and freshwater environments, which may be applicable to many elasmobranch species (see below). Despite the accomplishments of this dissertation, it is apparent that our understanding of the extra-renal mechanisms responsible for elasmobranch ion regulation and euryhalinity is far from complete and will continue to be an exciting area of future research. Below, I highlight a few of the remaining questions that I think would provide important contributions to the field of elasmobranch ion regulation if answered or addressed. What is the Function of the Rectal Gland in Freshwater Atlantic Stingrays? The results presented in Chapters 2 and 3 showed that rectal glands of freshwater Atlantic stingrays had a reduced size and lower Na+,K+-ATPase activity and expression, compared to rectal glands of marine Atlantic stingrays. This suggests that the gland has a lesser ability to secrete NaCl, but the fluid secreted from the rectal gland of a freshwater elasmobranch has never been collected. Therefore, the actual NaCl composition and secretion rate of the fluid are unknown. These would be critical parameters to measure in order to evaluate the function of the gland in a freshwater elasmobranch. Besides measuring rectal gland fluid output and composition, it would also be important to compare the expression of other components of the NaCl secretory pathway between freshwater and marine elasmobranch rectal glands. In this dissertation, I only examined Na+,K+-ATPase, but the Na+K+2Clcotransporter and cystic fibrosis transmembrane regulator (CFTR) Clchannel are also critical to NaCl secretion. As mentioned previously, CFTR is considered to be the rate-limiting step in active NaCl secretion (Riordan et al. 1994).

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119 What is the Mechanism of Branchial Na+/H+ Exchange? While I have shown that the V-H-ATPase-rich cells in Atlantic stingray gills are likely sites of Cl-/HCO3exchange via an apical pendrin-like transporter (see Chapter 5), I have not presented any data for an apical mechanism of branchial Na+ uptake and H+ excretion. However, Edwards et al. (2002) showed that apical Na+/H+ exchanger (NHE) isoforms are expressed in Na+,K+-ATPase-rich cells of marine elasmobranch gills. It is likely that similar NHEs exist in Na+,K+-ATPase-rich cells of seawater Atlantic stingrays to mediate Na+/H+ exchange, because a NHE would be driven by the favorable gradient for Na+ entry into the cells from seawater (Claiborne 1998, Claiborne et al. 1999). This gradient could be maintained by the basolateral Na+,K+-ATPase. In freshwater Atlantic stingrays, the mechanism responsible for branchial Na+/H+ exchange is unclear. One possibility is that freshwater stingrays use the same apical NHE isoforms found in marine elasmobranchs. Although an apical NHE driven by a basolateral Na+,K+-ATPase is not suitable for extremely low NaCl (< 0.5 mmol L-1) freshwater environments, an NHE could operate efficiently in a relatively high NaCl (> 1 mmol L-1) freshwater environment with an alkaline pH (Wright 1991). The fresh water of the St. Johns River and of the animal holding tanks used in my experiments had NaCl concentrations > 1 mmol L-1 and were alkaline; therefore, an apical NHE in Na+,K+-ATPase-rich cells is a possible mechanism for apical Na+/H+ exchange in freshwater stingrays. Future studies should attempt to identify NHE isoforms in the gills of freshwater and marine Atlantic stingrays to clarify this issue. How Can the Proposed Model of Branchial Ion Regulation Be Tested? In Chapter 5, I presented a model of NaCl and acid/base transport in the elasmobranch gill (Figure 5-9) that was based upon the biochemical and

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120 immunochemical findings of this dissertation and Edwards et al. (2002). To ensure that the proposed model is valid, it would be important for future studies to test this model and verify its components. An experimental approach to test if the V-H-ATPase-rich cells are responsible for Cl-/HCO3exchange would be to induce a metabolic alkalosis in freshwater or marine Atlantic stingrays and determine if the branchial V-H-ATPase-rich cells respond appropriately. For example, during an alkalosis I would expect to find an increase in the insertion of V-H-ATPase and pendrin into their respective plasma membranes to enhance HCO3secretion. To accurately measure these potential changes, it would be necessary to use immunocytochemistry, specifically immunogold localization followed by observations with a transmission electron microscope. With this technique it is possible to obtain high resolution images of the transporter localization that allow quantitative measurements of transporter expression in plasma membrane and cytoplasmic compartments. This approach has been used successfully to show that type B intercalated cells have more V-H-ATPase inserted into their basolateral membrane during a metabolic alkalosis (Verlander et al. 1992), which suggested the type B intercalated cells were important for HCO3secretion. What Controls Expression of Ion Regulatory Mechanisms In Chapters 3, 4, and 5, I showed that expression of Na+,K+-ATPase, V-H-ATPase, and a pendrin-like transporter in the gills of the Atlantic stingray are influenced by environmental salinity. I did not determine the endocrine mechanisms that control the salinity-induced changes in gene expression of these specific transporters, but it would be an interesting area of future research. The hormones that control gene expression of ion transporters in the elasmobranch gill have not been determined, but in teleosts, hormones such as cortisol, prolactin, and thyroid hormones influence branchial Na+,K+-ATPase

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121 activity, chloride cell proliferation, and chloride cell morphology (McCormick 1995). It would be valuable for future studies to determine if analogous hormones in the Atlantic stingray influence branchial ion transporter expression to identify possible endocrine mechanisms associated with ion balance in fresh and seawater environments. Is the Proposed Model Applicable to Other Elasmobranchs? In this dissertation, the Atlantic stingray was the only elasmobranch examined, which raises the question of whether the proposed model of branchial NaCl and acid/base transport is relevant to other elasmobranch species. The core of the proposed model is distinct populations of V-H-ATPase-rich and Na+,K+-ATPase-rich cells that are responsible for Cl-/HCO3and Na+/H+ exchange, respectively. By collecting gill tissue from species of various elasmobranch families and conducting the double-labeling immunohistochemical staining procedure conducted in Chapter 4, it can be determined if this model applies to other species. I have preliminary data that show distinct V-H-ATPase-rich and Na+,K+-ATPase-rich cells exist in the gill epithelium of marine elasmobranchs from the families Carcharhinidae, Lamnidae, Rajidae, Squalidae, and Gymnuridae, which suggests the proposed model of ion regulation is relevant to many elasmobranch species. It would be of great interest to determine if this model applies to the stenohaline freshwater Potamotrygonid stingrays of the Amazon River. This family (Potamotrygonidae) of stingrays evolved from a euryhaline ancestor that was closely related to Dasyatid stingrays (Lovejoy 1997), but have an osmoregulatory strategy that is similar to freshwater teleosts (e.g., lack of plasma urea, low plasma [NaCl]) (Thorson et al. 1967). Immunolocalization of V-H-ATPase and Na+,K+-ATPase in gills of Potamotrygonid stingrays would determine if this family’s relatively long isolation in

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122 fresh water has also resulted in a convergent pattern of V-H-ATPase and Na+,K+-ATPase expression with freshwater teleosts, or if the expression pattern more closely resembles the family’s Dasyatid ancestors. What Limits Elasmobranch Euryhalinity? In a review of the mechanisms involved with ion regulation by euryhaline fishes, Evans (1984b) listed, “What does limit elasmobranch euryhalinity?” as an important question to be answered by future researchers. Results from this dissertation suggest that euryhalinity of the Atlantic stingray is linked to this species ability to differentially express ion transporters in the gills (Na+,K+-ATPase, V-H-ATPase, pendrin) and rectal gland (Na+,K+-ATPase). For example, freshwater Atlantic stingrays were characterized by a greater expression of ion transporters associated with NaCl uptake and acid/base extrusion in their gills, compared to seawater individuals. Therefore, it is possible that one limit to elasmobranch euryhalinity may be the inability of a marine elasmobranch to increase expression of ion transport proteins in the gills during a low salinity encounter, which would decrease the animal’s ability to 1) extract NaCl required for osmotic regulation; and 2) excrete H+ and HCO3required for acid/base regulation. In addition, freshwater Atlantic stingrays were characterized by a lower rectal gland Na,K-ATPase activity and expression, compared to seawater individuals. Therefore, another possible limit to elasmobranch euryhalinity may be an inability of a marine elasmobranch to decrease rectal gland Na,K-ATPase activity and expression during low salinity exposure, which would increase losses of NaCl to the freshwater environment and complicate NaCl balance. It would be interesting to examine extra-renal ion transporter expression in elasmobranch species that are closely related to the Atlantic stingray, but have more

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123 restricted salinity tolerances (e.g., Dasyatis say, D. americana). This would provide an opportunity to test the hypothesis that elasmobranch euryhalinity is limited by the ability of a given species to differentially express ion transporters in their gills and/or rectal gland.

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127 Cutler, C. P., Brezillon, S., Bekir, S., Sanders, I. L., Hazon, N. and Cramb, G. (2000). Expression of a duplicate Na,K-ATPase beta(1)-isoform in the European eel (Anguilla anguilla). Am J Physiol 279, R222-229. Cutler, C. P., Sanders, I. L., Hazon, N. and Cramb, G. (1995). Primary sequence, tissue specificity and expression of the Na+,K+-ATPase alpha-1 subunit in the European eel (Anguilla anguilla). Comp Biochem Physiol 111B, 567-573. Dang, Z., Balm, P. H., Flik, G., Wendelaar Bonga, S. E. and Lock, R. A. (2000a). Cortisol increases Na+/K+-ATPase density in plasma membranes of gill chloride cells in the freshwater tilapia Oreochromis mossambicus. J Exp Biol 203, 2349-2355. Dang, Z., Lock, R. A. C., Flik, G. and Wendelaar Bonga, S. E. (2000b). Na+/K+-ATPase immunoreactivity in branchial chloride cells of Oreochromis mossambicus exposed to copper. J Exp Biol 203, 379-387. D'Cotta, H., Valotaire, C., Le Gac, F. and Prunet, P. (2000). Synthesis of gill Na+-K+-ATPase in Atlantic salmon smolts: differences in -mRNA and -protein levels. Am J Physiol 278, R101-110. De Vlaming, V. L. and Sage, M. (1973). Osmoregulation in the euryhaline elasmobranch, Dasyatis sabina. Comp Biochem Physiol 45A, 31-44. Degnan, K. J., Karnaky Jr., K. J. and Zadunaisky, J. A. (1977). Active chloride transport in the in vitro opercular skin of a teleost (Fundulus heteroclitus), a gill-like epithelium rich in chloride cells. J Physiol 271, 155-191. Degnan, K. J. and Zadunaisky, J. A. (1979). Open-circuit sodium and chloride fluxes across isolated opercular epithelia from the teleost Fundulus heteroclitus. J Physiol 294, 483-495. Degnan, K. J. and Zadunaisky, J. A. (1980). Passive sodium movements across the opercular epithelium: the paracellular shunt pathway and ionic conductance. J Membr Biol 55, 175-185. Dixon, T. E., Clausen, C., Coachman, D. and Lane, B. (1986). Proton transport and membrane shuttling in turtle bladder epithelium. J Membr Biol 94, 233-243. Donald, J. A., Toop, T. and Evans, D. H. (1997). Distribution and characterization of natriuretic peptide receptors in the gills of the spiny dogfish, Squalus acanthias. Gen Comp Endocrinol 106, 338-347. Edwards, S. L., Donald, J. A., Toop, T., Donowitz, M. and Tse, C.-M. (2002). Immunolocalisation of sodium/proton exchanger-like proteins in the gills of elasmobranchs. Comp Biochem Physiol 131A, 257-265.

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BIOGRAPHICAL SKETCH Peter Marc Piermarini was born in Mount Kisco, New York on May 8th, 1973 to Linda and Ernest Piermarini. He was raised in Cold Spring, NY, with his younger brother Craig, and went to Haldane Central School from kindergarten through high school. After graduating from Haldane in 1991, Peter attended James Madison University in Harrisonburg, Virginia, from 1991 to 1995 and received a Bachelor of Science in Biology, Summa cum Laude. During his studies at James Madison University, Peter also conducted an independent undergraduate research project under the supervision of Dr. Ann Pabst and received a Margaret Gordon Award for Outstanding Undergraduate Research in 1994 and 1995. In August of 1995, Peter started graduate school at the University of Florida, Department of Zoology working with Dr. David H. Evans. Peter chose to bypass his Masters degree in order to directly pursue a Ph.D. in the fall of 1997. During his studies at the University of Florida, Peter has been awarded seven research grants and fellowships, including an Environmental Protection Agency’s Science to Achieve Results Graduate Fellowship. Peter has also taught five different courses at the University of Florida, including a non-majors Biology lecture, and mentored seven undergraduate students who conducted independent research projects. On completion of his Ph.D., Peter plans to work as a postdoctoral scientist at Yale University, Department of Cell and Molecular Physiology, under the direction of Dr. Walter Boron. 146