Fine Structure, Innervation, and Functional Control of Avian Salt Glands

Fine Structure, Innervation, and Functional Control of Avian Salt Glands Rijdiger Gerstberger and David A. Gray Max-Planck-Institut fur Physiologische und Klinische Forschung, W. G. KerckhoffInstitut, D-6350 Bad Nauheim, Federal Republic of Germany

1. Introduction To maintain cellular functions, both the intracellular fluid volume and the composition of (non)-ionic solutes have to be controlled within narrow limits. Although single-celled organisms are separated from their mostly aqueous environment by only a plasma membrane endowed with specific transport systems, channels, and energy-driven pumps, multicellular organisms use their extracellular fluid space, with sodium and chloride as the major osmotically active ionic solutes, as a regulated exchange system between the intracellular compartment and the “external world.” In the vertebrate kingdom, various strategies to maintain body fluid homeostasis have evolved with regard to extracellular fluid volume (ECFV) and tonicity (ECIT), according to the axiom of Bernard (1865), which is still a central tenet of comparative physiology. With the kidney and its transport systems representing the main osmoregulatory organ in mammalian salt and water balance, accessory tissues highly specialized for the transport of sodium, chloride, or divalent ions in combination with osmotically induced water fluxes are described for submammalian vertebrates and include the gills of teleost fish (Foskett, 1987), the rectal gland of elasmobranchs (Solomon er a/., 1984a, 1985a), the skin of amphibians (Lindemann and Voute, 1977), the gut system of fish and birds (Skadhauge, 1981; Kirsch et al., 1985), and the salt-secreting glands of marine reptiles and birds (Peaker and Linzell, 1975; Dunson, 1976). The unusual position of the avian salt-secreting glands in the head region of marine birds and their accessibility have stimulated scientific examination in the past with first reports appearing as early as 1665 (Technau, 1936). With detailed descriptions of the exact anatomical localization of these glands including their duct systems in various families of the bird kingdom, Jacobson (1813) and Inr~rnorronalRewen, OJ Cyrologr. Vol. 144

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Nitzsch (1820) summarized the knowledge of their time without presenting information concerning the fine structural aspects of the glands, or any indication of the putative physiologic function of the glands. Nitzsch only stated that “the possibly oily nature of the fluid can be excluded as it quickly evaporates when soaked up by paper” and that “the secreted fluid might resemble drops of tears.” Ontogenetic studies concerning the development of glandular structures in the head region of various vertebrate classes then created the false idea that the salt gland was homologous to the organ of Jacobs, involved in chemoreception (Kolliker, 1860; Mihalkovics, 1898). It was not until the early twentieth century that Cohn (1903) managed to rule out the participation of the salt gland in sensory processes. Numerous observations then described the close relation between the size of the salt glands and the salinity of the water in the respective habitat for a variety of primarily marine, but also terrestrial, avian species (Marples, 1932). Fast increases in the specific organ weight of the salt glands in ducklings (Anus platyrhynchos) adapted to 3% saltwater as drinking fluid as well as the reduction of salt gland size in Eider ducks (Somateria rnoflissima) kept on freshwater strongly supported this relationship (Heinroth and Heinroth, 1928). More than 100 years after the pioneering studies by Nitzsch and Jacobson, Technau (1936) published a detailed analysis of the avian salt-secreting gland, emphasizing the whole spectrum of exact localization, macromorphology, and formation of the duct system of salt-secreting glands obtained from 106 genera of birds, including such rare species as Urubu, South American Phytotoma, and islandic anatides. With regard to the physiology of the glands, however, Technau (1936) only dedicated a few words to its function, supporting the idea of Heinroth and Heinroth (1928) and Schildmacher (1932), who reported that the salt glands of the Eider duck secrete a fluid that “protects the nasal mucosa against irritating influences of the salty seawater.” The often reported observation of fluid droplets at the tip of the beak during open-sea feeding of truly marine birds such as petrels, albatrosses, and penguins (Peaker and Linzell, 1975), and the urgent need to eliminate the excessively ingested salt from their extracellular fluid compartment at the restricted concentrating capacity of their kidneys, finally led to the discovery of the important role that avian salt glands play in maintaining body fluid homeostasis (Schmidt-Nielsen et al., 1958; Schmidt-Nielsen, 1960).

II. Secretory Tissue of the Avian Salt Gland A. Zoology of Salt Glands The gross anatomy, fine structure, secretory mechanism, and hormonal control of the avian salt-secreting gland have been thoroughly revised over the last two

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decades (Peaker and Linzell, 1975; Van Lennep and Young, 1979; Butler, 1984; Holmes and Phillips, 1985; Komnick, 1985; Butler et al., 1989, Simon and Gray, 1989; Gerstberger, 1992), following the pioneering work of SchmidtNielsen (1960). In the present review on various aspects of fine structure, innervation, and blood supply, and also afferent and efferent control mechanisms of avian salt gland function, new insights gained within the last 10 years are incorporated with well-known observations that now can be looked upon from a different point of view. These recent findings should shed light on (a) the functional cellular architecture, ( 6 )the osmo- and volume-receptive system involved in salt gland control, (c) the efferent effectors such as neurotransmitters and hormones, and (d) the intracellular signal transduction pathways. Functional salt glands have been demonstrated in at least 10 avian orders including Charadriiformes (gulls and plovers), Procellariiformes (albatrosses, petrels) (Fig. I), Pelicaniformes (pelicans, cormorants), Sphenisciformes (penguins), Gaviiformes (divers), and Anseriformes (ducks, geese), as well as Falconiformes (eagles) and birds living in arid land zones (Peaker and Linzell, 1975; Thomas and Phillips, 1978; Mahoney and Jehl, 1985; Sonawane, 1987; Conway et al., 1988). The exact location of the avian salt-secreting gland in the head region differs depending on the species and/or its habitat. The salt glands of wading birds, plovers, gulls, and petrels are found in the crescent-shaped depressions in the frontal bones above the eyes (Fig. 8), whereas the salt glands of ducks and geese are located along the edge of the frontal bone in the upper orbital membrane. The salt glands of falcons and woodpeckers are in the upper maxillary cavity and beneath the orbital membrane, respectively. Concerning the embryological origin of the salt glands, Marples (1932) stated that the gland is eventually formed by branching of the two main ducts arising from the rudiment of the nasal cavity, and then growing backward to the final position. Histological studies performed with the Adelie penguin (Pygoscelis adeliae) allowed the first rudiments of supraorbital salt glands to be traced back to “solid crescent structures on either side of the nasal cartilage,” growing posteriorly to develop dorsal to the eye (Herbert, 1975). Thus, the glandular matrix develops from the ducts as branched tubules radiating from a central canal. Salt-excreting glands evolved in marine birds to eliminate excess ions, mainly sodium and chloride, from the extracellular fluid compartment to maintain body fluid homeostasis even under severe osmotic stress (Bemdge and Oschman, 1972; Kirschner, 1980). Reptiles living in marine, estuarine, and arid zones and marine elasmobranchs also developed “extrarenal glandular organs whose main function is to excrete a hypertonic salt solution” (Van Lennep and Young, 1979). The salt-secreting glands of reptiles comprise lingual glands in crocodilians, lacrimal glands in chelonians, sublingual glands in snakes, and lateral nasal glands in lizards, whereas the nonhomologous shark rectal gland represents a rectal appendix in the dorsal mesenteries of the peritoneal cavity (Bulger, 1963; Dunson, 1976; Komnick, 1985). Whenever appropriate, comparative aspects of

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FIG. 1 Marine birds such as this adult New Zealand albatross possess supraorbitally located saltsecreting glands (salt glands), with their main ducts opening into the nasal cavity, to maintain body fluid homeostasis in their salty environment.

salt-secreting gland structure and function in elasmobranchs or reptiles have been taken into consideration in this review. The salt glands of marine and estuarine birds, or of saltwater-acclimated freshwater birds bearing salt glands, are able to actively extract sodium and chloride from the extracellular fluid compartment, represented by the perfusing bloodstream, and to excrete up to 1.O ml/min/g gland weight of a hypertonic salt solution against a marked concentration gradient. Sodium is accompanied by chloride in equimolal concentrations, whereas potassium concentrations in the secreted fluid remain low. Depending on the species, sodium concentrations range from about 500 meq/liter, as described for the saltwater-acclimated duck or some cormorants, to more than 1200 meq/liter, as reported for petrels and albatrosses (Schmidt-Nielsen, 1960; Peaker and Linzell, 1975). Both sodium and chloride therefore must be actively transported against a marked concentration gradient of 1:3 to 1 :8, compared to the mere ratio of 1:1.5 for the shark rectal gland. Like the avian salt gland, most reptilian salt glands are able to highly concentrate sodium and chloride from the perfusing bloodstream, thereby elim-

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inating excess salt from body fluids at low water losses. In herbivorous lizards, potassium and bicarbonate constitute a major portion of the excreted electrolytes (Dunson, 1976; Van Lennep and Young, 1979). In many exocrine glands, the rate of secretion is closely correlated to the concentration of solutes in the secreted fluid. For the avian salt gland, however, this aspect has been discussed controversially. With salt gland secretion rates in the duck varying from 0.2 to 0.8 ml/min, there appeared to be no correlation between secretion rate and sodium concentration of the fluid with ion concentrations being at their maximal level (Butler et al., 1989). At secretion rates lower than 0.1 ml/min, however, the concentrations of sodium and chloride were concomitantly reduced (Deutsch et al., 1979; Gerstberger et al., 1984a). A comparable relationship was reported for the Kelp gull (Gray and Erasmus, 1989b). In the goose, either a slightly positive or a negative correlation was observed, depending on the salt and volume loading of the animals (Hanwell et al., 1971a).

6 . Fine Structure of the Secretory Cell The avian salt gland represents a compound tubular gland, with its main excretory ducts branching into primary and secondary ducts forming the central canals of secretory lobes (or lobules) (see Section II,D), which are composed of numerous radially arranged secretory tubules with small lumina merging into these central canals (Van Lennep and Young, 1979) (Fig. 2). Different from the salt glands of the gulls, plovers, or the oyster catcher, where secretory lobes running parallel all drain into one of the two main ducts leaving the secretory tissue, the duck’s salt gland is composed of spherical secretory lobes. Central canals of these lobes, receiving inflow from all the lumina of radiating secretory tubules, empty into primary ducts, which finally merge with the medial or lateral segmental ducts (Butler et al., 1991). Sheaths of peritubular connective tissue and a dense interlobular connective tissue matrix containing nerve fibers and blood vessels embed single secretory tubules and lobes, respectively. The first detailed morphologic description of the avian salt gland at the electronmicroscopic level was camed out by Doyle (1960) using specimens of the Great Black-Backed gull (Larus marinus) and the petrel (Oceanodroma leucorrhoa). Subsequently, the salt glands of L. argentatus, L. ridibundus, and the saltwateracclimated Pekin duck (A. platyrhynchos) were subjected to basic light- and electron-microscopic examination (Komnick, 1963a,b,c, 1964; Dulzetto, 1965; Ernst and Ellis, 1969). 1. The Principal Cell

The typical secretory tubule of the avian salt gland comprises five to eight cells radially arranged around a central lumen of 1-1.5 pm in diameter, thus forming

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a monolayered epithelial sheath (Fig. 2 ) . The secretory tubules radiating from each central canal are lined by a cuboidalxolumnar epithelium of principal secretory cells at the proximal, and so-called terminal or peripheral cells at the blind-ending distal segments (Emst and Ellis, 1969). Morphometric analysis allowed the total surface area of the secretory tubule lumina to be determined as 3.65 cm2 in the saltwater-acclimated Pekin duck (Marshall et al., 1987). The peripheral cells revealing rather high mitotic activity (Fig. 5) are characterized by simple, unspecialized cytoarchitecture with high numbers of unbound ribosomes, but low density of basally located mitochondria. The basolateral membranes appear unfolded with only a few plicae detectable. Furthermore, the cells are equipped with large Golgi apparatus (Ernst and Ellis, 1969). The fully specialized principal cells constituting the major portion of the secretory tubules in salt-acclimated animals (Fig. 5 ) possess a large spheroidal nucleus located in the center of the cell or slightly above it. These cells reveal maximal surface membrane amplification due to marked interdigitations at the basal, but also lateral, cell borders (Figs. 2 and 3). The basal lamina embedding a secretory tubule remains straight and does not follow the plasma membrane invaginations that enwrap a great number of mitochondria (Komnick, 1963a,c; Emst and Ellis, 1969). These mitochondria are of elongated shape, characterized by densely packed cristae within their inner matrix, and situated within the basal infoldings of the plasma membrane (Komnick, 1963c) (Fig. 3). Purified mitochondria1 preparations enabled the calculation of ADP utilization per molecule cytochrome c, as well as of sodium transport with maximal values of eight sodium ions (Na+) per molecule ATP (Chance et al., 1964). “Secretion canaliculi,” once proposed by Doyle (1960), are fully absent from the electrolytetransporting epithelium, which is composed of the avian principal secretory cells (Van Lennep and Young, 1979). Different from the basolateral membrane, the apical border shows only small microvilli protruding into the small lumen of the secretory tubule. Adjacent principal cells are apically interconnected via junctional complexes comprising zonulae occludentes, zonulae adherentes, and desmosomes (Fig. 4). To outline the basolateral membrane surface with all its plicae, interdigitations, and complex foldings, horseradish peroxidase, ruthenium red, and lanthanum salts were used

FIG. 2 The secretory parenchyma of the functional avian salt gland. (a) Light-microscopic cross section through a lobe of an Eider duck salt gland (Masson Goldner staining), with secretory tubules (ST) radially arranged around the central canal (CC). Bar, 30 pm. (b) Light-microscopic cross section through secretory tubules (ST) of an Eider duck salt gland (Azan staining) with five to eight principal cells surrounding the tubular lumen (L). Countercurrent arrangement of secretory tubules and capillaries (CA). Bar, 10 pm. (c) Electron micrograph of a cross section through one secretory tubule of a Pekin duck salt gland revealing intercellular luminal tight junctions (TJ), basolateral infoldings (IN), and nuclei (N) of seven secretory cells. Bar, 1 pm. (Unpublished micrographs, courtesy of Prof. Dr. W. Kiihnel.)

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FIG. 3 The basolateral plasma membranes of avian salt gland secretory cells. (a) Extensive basolateral infoldings (IN) of a salt gland secretory cell (Pekin duck) as demonstrated by lanthanum treatment. BL, basal lamina; N, cell nucleus; L, lumen. Bar, 1 pm. (b) Basal membrane infoldings (IN) of a salt gland secretory cell (Japanese swan goose) with numerous cristae-type mitochondria (M). The base of the secretory cells is in contact with a continuous basal lamina (BL). Bar, 1 pm. (c) Freeze-fracture replica of the basal infoldings from a secretory cell of a saltwater-acclimated Pekin duck. Mitochondria (M) are densely packed within the intracellular compartments. BL, basal lamina; P, particle-rich fracture faces; E, particle-poor fracture faces. Bar, 0.5 pm. [(a) From Berridge and Oschman. 1972, with permission; (b) Unpublished material, courtesy of Prof. Dr. W. Kiihnel; (c) from Riddle and Ernst, 1979, with permission.]

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to mark the extracellular interstitial space between neighboring secretory cells in saltwater-acclimated ducklings (Martin and Philpott, 1973, 1974; Hootman et al., 1987) (Fig. 3). Using these extracellular space tracers, both the extent of the space and the permeability of the junctional complex could be examined in actively secreting salt glands. Ruthenium red reacts with anionic components of the cell surface. Horseradish peroxidase could be found filling the matrix area of the peritubular connective tissue and the spaces formed by the basolateral cell membrane infoldings and in the junctional complexes and even the luminal space. Penetration into the lumen of the secretory tubules was also described for lanthanum, suggestive of rather leaky cell-to-cell contacts (Hootman et d., 1987).

FIG. 4 The luminal junctional complex in the avian salt gland secretory tubule. (a) Narrow lumen (L) of a secretory tubule from a Pekin duck salt gland bounded by six secretory cells joined lumiSurface specializations such as short microvilli (MV)protrude into the nally by tight junctions (TI). lumen. Bar. 0.5 pm. (b) High magnification of the junctional complex between adjacent principal cells in the Herring gull salt gland. The intercellular gap (I) is bridged by a zonula occludens ( O ) , zonula adherens (A), and desmosome (D). Bar, 0.1 pm. (c) Freeze-fracture replica of the zonula occludens of secretory cells from a Pekin duck salt gland. A junctional douplet consisting of two strands is visible with the fracture running from the P face (PF) of the luminal membrane to the P face (P) of the lateral membrane. The two-stranded junction can be locally widened (80 nm) with interconnecting cross bars (asterisk). Bar. 0.1 pm. [(a) from Ernst and Ellis, 1969, reproduced with permission from J . Cell.. B i d . (1979). 40. 231-305; (b) from Komnick and Kniprath, 1970, with permission; (c) from Riddle and Ernst, 1979, with permission.]

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To elucidate cellular mechanisms involved in the ion transport processes and their control systems, the intact avian salt gland proved to be of limited use, whereas the isolated secretory cell represents an ideal model. Barmett and colleagues (1983), however, opposed the use of long-term cell cultures of salt gland secretory cells due to the dedifferentiation including the loss of polarity and basolateral infoldings. Some of the latter problems could possibly be overcome using confluent sheets of dissociated cells seeded on a porous collagen support that expressed polarized features such as apical microvilli, tight junctional complexes with transmural resistance, and basolateral membrane infoldings (Lowy et al., 1985a,b) (Fig. 14). Enzymatic and mechanical dissociation of salt gland tissue gave high yields of viable principal cells readily identified in suspension by their large size, cytosomal granularity, and retained cell surface plicae. Apical polarity of these cells might, however, be lost (Hootman and Emst, 1980). 2. Apical Tight Junctions As mentioned, leakiness of the secretory epithelium was indicated by extracellular space markers and also the retrograde injection of India ink or colloidal thorium via the excretory duct of the gland. This procedure resulted in penetration of the ink from the tubular lumen to the basement membrane (Doyle, 1960). By transmission electron microscopy (EM) of the apical junctional complex, the zonula occludens in the secretory cells of the avian salt gland, possibly proteinaceous in nature, appeared to be a component of the epithelial junctional complex, which includes the zonula adherens (intermediate junction) and the desmosomes (Fig. 4). In freeze-fracture EM, the tight junction appeared as continuous, intramembrane strands in the P-face (outwardly facing cytoplasmic leaflet) with complementary grooves in the E-face (inwardly facing extracytoplasmic leaflet). In general, tightness of an epithelium is represented by the correlation of freeze-fracture strand number and transepithelial resistance (Claude and Goodenough, 1973). Freeze-fracture studies in the nasal salt gland of the Herring gull demonstrated apical cell junctions of the single-stranded type (Ellis et al., 1977). Freezefracture replicas of principal secretory cells of the duck salt gland, obtained from the particle-rich (P) face of the luminal membrane to the particle-rich (P) face of the basolateral membrane, also revealed shallow luminal intercellular junctional complexes consisting of only two closely juxtaposed junctional strands on the P face of the lateral membrane. With the exception of short focal widenings, two sets of doublets were rarely seen, each doublet having a width of 20-25 nm (Riddle and Emst, 1979) (Fig. 4). The simplicity of the junctional morphology in the avian salt gland indicates leaking of ions and other solutes, quite in contrast to the complex network of anastomosing strands, described for high-resistance epithelia such as frog skin or endothelial cells forming the blood-brain banier

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(Claude and Goodenough, 1973; Simons and Fuller, 1985; Gumbiner, 1987). The peripheral, transport-inactive cells possess zonulae occludentes composed of a loose meshwork of interconnecting strands at a junctional width of about 100 nm. Interestingly, principal cells from salt glands of osmotically unstressed ducks showed zonulae occludentes of slightly higher complexity, the strands often being made up of two sets of doublets (Riddle and Emst, 1979). Although direct access to the glandular tubules has not been possible in vivo, transepithelial resistance can be used as a measure for tightness of junctional complexes between cells, determined from confluent layers of primary salt gland cell culture systems (Lowy et al., 1985b) (Fig. 14). Values in the range 300 R X cm2 represent only modest resistance compared to those obtained from brain endothelial cells or frog skin (2000 R X cm2) (Bradbury, 1985).

3. Comparative Aspects Reptilian salt glands such as those found in the desert lizard Uromastyx acanthiniirus are composed of two types of electrolyte-transporting principal cells, called “light” and “dark” cells according to the electron density of their cytoplasm, mainly due to the packing of large numbers of mitochondria. Whereas the light cells are ovoid shape, the dark cells are triangular in shape. The apical membranes of the principal cells of the salt gland reveal short microvilli, as in the avian salt gland, except for a few cells endowed with apical brush borders (tuft cells), whereas the lateral membranes are strongly plicated (Van Lennep and Young, 1979). In addition, cells containing a well-developed rough endoplasmic reticulum and secretion granules in the apical zone, stained with periodic acid-Schiff’s reagent (mucoid cells), and so-called “basal cells” because of their location at the base of the secretory tubules have been described (Van Lennep and Komnick, 1970). The posterior sublingual salt glands of all sea snakes investigated so far lie beneath the tongue in the lower jaw and partially enclose the tongue sheath, into which they secrete. The ultrastructure of the sublingual salt gland in sea snakes resembles that of other reptilian salt glands with their highly irregular principal cells being rich in mitochondria and plasma membrane (baso-) lateral digitations. Quite different, however, adjacent apical cell surfaces appear to be joined by zonulae adherentes and not occludentes, implying a continuity between the interstitial space and the tubular lumen. Systemic salt loading in the poisonous pelagic sea snake Pelumis induced secretion from its large single posterior sublingual salt gland with sodium (“a+]) and chloride ([Cl-]) concentrations as high as 700 mM at low potassium ([K+]) concentrations of 15 mM, results comparable to those observed with the avian salt gland (Dunson and Taub, 1967; Dunson, 1968, 1976; Dunson et al., 1971; Dunson and Dunson, 1974). The dogfish rectal gland is composed of a central canal surrounded by an inner layer of transitional epithelium, a broad layer containing the secretory

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tubules, and the outer capsule. The principal cells of the secretory tubules, described in detail by Bulger (1963), reveal pronounced membrane interdigitations at the basal as well as lateral cell borders. Comparable to the avian secretory cell, mitochondria of the cristae-type are densely packed in the basal cellular compartment, and endoplasmic reticulum is found only in small amounts. The Golgi apparatus can be recognized as a separate unit close to the nuclear membrane. Comparable to the shallowness of the junctional complex between neighboring salt gland cells in birds, occluding junctions between secretory cells of shark and stingray rectal glands showed an average strand nurnber of 2 and 3.5, respectively. Apical boundaries of the cells are quite tortuous, giving rise to a long linear junction and large paracellular conductance/unit apical surface area (66 m/cm2). The measurements are similar to those obtained for the avian salt gland (S. A. Emst, personal communication). As emphasized by electrophysiologic studies, these epithelia might also be of the leaky-type (Emst et al., 1981; Greger et al., 1986).

C. Cellular Aspects of Salt Acclimation 1. Glandular Hypertrophy To become functionally active, the salt glands of newly fledged marine or young freshwater-acclimated birds must undergo a complex adaptive process, finally leading to the fine structural aspects described for the secretory principal cell. The most obvious signs of salt acclimation are represented by increased organ weight and augmented secretory capacity. This could be described for adult Pekin ducks, Australian chestnut tails (A. castanea), and Glaucous-winged gulls (L. glaucescens) during long-term salt acclimation (Holmes et al., 1961; Ellis et al., 1963; Fletcher et al., 1967; Deutsch et al., 1979; Baudinette et al., 1982). Acclimation of newly hatched common gulls (L. canus), Glaucous-winged gulls, or ducklings to saline in their food supply also induced glandular hypertrophy (Schmidt-Nielsen and Kim, 1964; Spannhof and Jiirss, 1967; Gerstberger et al., 1984a; Hughes, 1984) (Fig. 5). Deadaptational processes resulted in opposite responses. Quantitative morphometric analysis of structural differences between salt glands of Hemng gulls acclimated to freshwater compared to those maintained on seawater yielded a decrease in whole organ size to 60% of the control values, with the intercellular fluid volume and the number of cytosolic mitochondria both diminished by half (Komnick and Kniprath, 1970). Deadaptation also resulted in a fast decline of Na-K-ATPase activity (see Section V,A), movement of the mitochondria toward the cell nucleus, and, importantly, loss of plicated plasma membranes. Clusters of filamentous osmiophilic material were found in the apical cytoplasm, and both acid phosphatase and peptidase activities were stimulated (Hossler et al., 1978).

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Variations in size and function of the salt glands have been reported for Franklin’s gulls (L. pipixcan) breeding in freshwater habitats and wintering in marine habitats (Burger and Gochfield, 1984), and for female Eider ducks (S. mollissima) during egg incubation, lasting for 26 days, when not drinking saltwater. The nasal salt glands concomitantly underwent regression and reduction in Na-K-ATPase activity and secretory capacity, thus representing a naturally occumng phenomenon of deadaptation (McArthur and Gorman, 1978). Another important aspect of salt acclimation and its role in avian salt gland function is the occurrence of alterations in body fluid distribution (see Section VI). Studies have been restricted to some gulls species, Canada geese, and the Pekin duck (Roberts and Hughes, 1984; Gray et al., 1987; Brummermann. 1988; Gray and Erasmus, 1989b; Hughes, 1989; Brummermann and Simon, 1990). During the course of salt acclimation, plasma electrolyte concentrations of Pekin ducks increased at constant blood volume, whereas the apparent volume for inulin distribution diminished, reflecting a reduced extracellular, especially interstitial, fluid volume (Gray et al., 1987). With regard to the size of the extracellular fluid compartment, however, data have been conflicting with bromide spaces identical for adapted and nonadapted birds (Ruch and Hughes, 1975), and with exchangeable sodium pool size being either of equal value (Gray et al., 1987) or increased after salt acclimation (Roberts and Hughes, 1984; Hughes, 1989). In the gull L . glaucescens, saline drinking did not influence the sodium pool size (Roberts and Hughes, 1984), whereas the gradual adaptation of Kelp gulls to 100% seawater as the drinking fluid resulted in a change of blood hematocrit indicative of extracellular fluid depletion, with a mild increase in plasma electrolyte concentrations but unchanged colloid osmotic pressure (Gray and Erasmus, 1989b).

2. Cellular Hypertrophy/Hyperplasia and Membrane Biogenesis Hypertrophy and hyperplasia of principal secretory cells during salt acclimation have been shown in purely morphometric studies performed on Pekin ducks, where short-term acclimation to a hypertonic saline regimen caused an increased average cell diameter of 11.6 pm compared to 10.4 pm under control conditions (Ballantyne and Wood, 1969). Cell densities per unit of tissue volume were reduced by half (Merchant et al., 1985). A fivefold increase in mitochondrial profiles and a sevenfold rise in the number of lipid droplets were also described as a result of acclimation to a hypertonic saline regimen (Merchant et al., 1985) (Fig. 13). In addition, enhanced cellular protein synthesis (Holmes and Stewart, 1968), increased cellular proteinDNA ratios (Hootman and Emst, 198la; Hossler, 1982), the ribosomal composition of principal secretory cells including augmented relative RNA content of the cell (Holmes and Stewart, 1968; Ballantyne and Wood, 1969; Stewart and Holmes, 1970; Knight and Peaker, 1979), and the stimulated incorporation of tritiated thymidine into

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FIG. 5 Morphologic indications of adaptation to a high salt regimen. (a) Schematic drawing showing the effects of a saltwater regimen on the development of avian salt gland secretory cells, and their distribution along a single secretory tubule. Partially specialized cells (Day 0, stippled) become endowed with basolateral infoldings and numerous mitochondria (Day 2, hatched), and develop pronounced basolateral membrane amplification and cellular polarity (Day 1I , cross-hatched). Fully matured principal secretory cells (hatched) are found near the merging point of the branched secretory tubule with the central canal. (b) Cross section through the salt gland of a freshwater-adapted (1) and a saltwater-adapted (2) Pekin duck (Azan staining). Bar, 1 mm. (c) Histochemical demonstration of acyltransferase activity as a marker enzyme of membrane biogenesis in salt gland secre-

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nuclear DNA (Knight and Peaker, 1979) were used as representative measures of cellular hyperplasia and hypertrophy. Geese with unilateral postganglionic denervation or surgical removal of one of their salt glands responded to shortterm salt acclimation with partially compensatory growth of the remaining gland, as judged from increased RNA concentrations per unit mass of tissue and an augmented RNADNA ratio in the intact gland (Hanwell and Peaker, 1975; Knight and Peaker, 1979). As suggested by the studies of Emst and Ellis (1969), the hypertrophic process commences at the blind end of the secretory tubules, where peripheral cells become mitotically active, as indicated by the 15-fold stimulated incorporation of radioactive thymidine after 1 day of salt acclimation. During the first hours of salt stress, radiolabeled thymidine was preferentially taken up by connective tissue cells, whereas at later stages, the label was concentrated in parenchymal cells of peripheral tubular origin (Hossler, 1982) (Fig. 5). These newly created cells would then transform to partially and finally fully specialized principal secretory cells (Fig. 5 ) . Plasma membrane biosynthesis in the salt gland secretory cells was markedly enhanced during salt acclimation of ducklings. Elucidation of the development of the fine structure of the secretory epithelium during salt acclimation at the ultrastructural level was facilitated by the marked amplification of the plasma membrane (Emst and Ellis, 1969; Hossler et al., 1978). The principal secretory cells of the avian salt gland revealed pronounced alterations in the basal and lateral plasma membrane with values of 1800, 9500, and 3500 pm2/cell before, during, and after saltwater acclimation in ducks (Fig. 13). In contrast, there were no detectable differences in the membrane surface area of the peripheral cells (Merchant et al., 1985). Stimulated synthesis of RNA and (g1yco)-proteins could be detected as early as 2 hr after the beginning of the salt stress, as determined from the incorporation of tritiated uridine, leucine, or fucose into salt gland tissue (Sarras et al., 1985). Pulse-chase experiments at the electronmicroscopic level indicated that [3H] leucine is initially taken up by the rough endoplasmic reticulum, transported to and concentrated in the Golgi apparatus, and finally found throughout the plasma membrane, with the three steps traceable after 5 , 10, and 60 min, respectively.

tory cells during saltwater acclimation is confined to some cisternae of the Golgi apparatus (G). Bar. 0.5 pm. (d) Autoradiograms of a salt gland secretory lobule of a Pekin duck salt-acclimated for 6 days following [3H]thymidine incorporation. In the darktield micrograph (top), labeled nuclei are indicated by double arrowheads, unlabeled nuclei by single arrowheads. In the lightfield micrograph (bottom), labeled parenchymal cells are indicated by single arrowheads, labeled connective tissue (CT) cells by double arrowheads. Bar, 20 pm. [(a) From Ernst and Ellis, 1969 (modified), with permission; (b) from Gerstberger e r a / . , 1984a. with permission of Springer Press, Heidelberg; (c) from Barmett e r a / . , 1983, with permission; (d) from Hossler, 1982. with permission.]

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Plasma membrane phospholipids increased by a factor of 1.7 during enhanced membrane biogenesis associated with cellular salt acclimation, and phospholipids demonstrated by Sudan Black or thionine were highly concentrated in the principal, but not peripheral, cells (Ellis et al., 1963). Acyl transferase activity, used as marker of acyl lipid biosynthesis, increased almost threefold during the first 24 hr of salt acclimation. The cytochemical localization of acyl transferase activity was restricted to some cisternal elements of the Golgi apparatus, indicative of phospholipid synthesis in this organelle of the principal cells with subsequent packaging and vesicular transport to the preexisting plasma membrane (Levine et al., 1972; Barrnett et al., 1983) (Fig. 5). Acid phosphatase, another enzyme possibly involved in membrane turnover, was found throughout the secretory tubules except in their peripheral blind-ending section, with highest densities of the enzyme in the basolateral region of the principal cells, which suggests local membrane degradation (Ellis et al., 1963; Spannhof and Jurss, 1967; Hossler et al., 1978). In contrast, alkaline phosphatase was moderately concentrated in the basophilic peripheral cells and glandular capillaries, but virtually absent from principal cells (Scothorne, 1958, 1960; Ellis et al., 1963; Spannhof and Jurss, 1967). As suggested, alterations in salt gland ultrastructure have often been associated with the osmoregulatory status of the animal. The volume fraction of some cellular organelles, such as mitochondria, and the size of the intracellular space have been used to draw far-reaching conclusions on “physiologic significance.” A detailed stereological analysis of the effects of buffer osmolality alone on the salt gland epithelium during processing of the tissue strongly questions whether the changes in intercellular space geometry reported “represent physiologic mechanisms in vivo or differences in osmotic behavior during processing for electron microscopy” (Cowan, 1986). Morphometric analysis of cellular structures thought to be most relevant for the secretory process, the mitochondria and the intercellular spaces, should therefore only be performed under standardized conditions of tissue perfusion and fixation, as indicated by Butt and colleagues (1985).

3. Metabolic Enzyme Activities Another important aspect of both salt acclimation of the principal cells and their energy-consuming transport activities is presented by the specific activities of various soluble enzymes involved in energy metabolism. When various soluble enzymes in the salt glands of the gull and Pekin duck were measured, those involved in cellular metabolic processes such as mitochondria1 respiration, glycolysis, and the Krebs cycle appeared to supply most of the energy (A”) needed for the ion transport activity (McFarland el al., 1965; Kiihnel et al., 1969a,b; Stainer et al., 1970). In addition, in vitro studies revealed that the presence of Krebs cycle substrates greatly increased cellular respiration (Borut and Schmidt-Nielsen, 1963).

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Acclimation to hypertonic saline as drinking water in the Pekin duck stimulated the specific activities of soluble enzymes involved in glycolysis and the hexose-monophosphate shunt such as glucose-6-phosphate dehydrogenase (G6-PD) and phosphofructokinase (PFK), which is often regarded as the ratelimiting enzyme in glycolysis. Hexokinase (HK) activity was slightly stimulated and lactic dehydrogenase (LDH) remained unchanged (Stainer et al., 1970). Enzyme analysis performed in salt gland tissue of saltwater-acclimated western gulls (L. occidentalis) resulted in high activities of G-6-PD and LDH, indicative of the importance of the hexose monophosphate shunt and rapid anaerobic glycolysis (McFarland et al., 1965). With regard to enzymes of the Krebs cycle, in the duck salt gland isocitric dehydrogenase (ICD) and malic enzyme (ME) activities remained unchanged during salt acclimation, whereas succinate dehydrogenase (SDH) activity was markedly enhanced (Ellis et al., 1963; Stainer et al., 1970). Succinate dehydrogenase activity also increased during the first days of salt acclimation in both proximal and distal components of the salt gland secretory tubules in young common gulls, whereas in nonadapted hatchlings the enzyme was restricted to the proximal aspects (Spannhof and Jurss, 1967). The moderate (duck) to very high (gull) activity of glutamic-oxaloacetic transaminase (GOT) and the low activity of glutamic-pyruvic transaminase (GPT) in the gull salt gland might be indicative of the potential use of the glutamate-aspartate shunt as possible source of energy (McFarland et al., 1965; Stainer el al., 1970). Adjusted calculations of energy yields from glycolysis and the Krebs cycle including the hexose monophosphate shunt were in the order of 300% of the energy needed for the secretory process.

D. The Duct System 1. Avian Salt Glands Jobert (1869) was the first to describe the avian salt gland as being formed of two distinct segments with separate drainage ducts. The putative role of these ducts in the modification of the transepithelially secreted fluid remains unclear to date (Marshall et al., 1985). In gulls and other Charadriiformes investigated, the secretory lobes running parallel all empty via their central canals directly into one of the two ducts leaving the salt gland at its anterior edge (Fange et al., 1958a,b). In ducks, the salt gland consists of distinct medial and lateral segments, each comprising spherical secretory lobes. The external duct of the medial segment empties into the nasal cavity at the base of the vestibular fold; the external duct of the lateral segment opens onto the surface of the nasal septum (Fig. 6). As revealed by microfil injections into both drainage ducts, the medial segment covers two-thirds of the salt gland (Butler et al., 1991) (Fig. 6). Also different from the situation in the gull, anastomoses between the lateral and the medial segments of

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FIG. 6 The duct system of the avian salt gland. (a) Schematic drawing of the nerve supply to the right nasal salt-secreting gland of a saltwater-acclimated Pekin duck (left), and the location of its medial and lateral drainage ducts in the nasal cavity (right). (b) Transverse section of a Pekin duck salt gland with medial (dark staining) and lateral (light staining) segments filled by retrograde injection of colored microfil into the medial (MD)and lateral (LD)duct. PD,primary duct merging into MD. Bar, 500 pm. (c) Lining of the medial duct (Azan staining) in an Eider duck salt gland. Bar, 30 pm. (d) Scanning electron micrograph of the columnar cells lining the medial duct of a Pekin duck salt gland. Small microvillar protrusions face the lumen of the duct. Bar, I pm. [(a,b) From Butler ef (11.. 1991; copyright 0 1991, reprinted by permission of Wiley-Liss, a division of John Wiley and Sons, Inc. (c) Unpublished material, courtesy of Prof. Dr.W. Kiihnel.]

the salt gland duct system appear not to occur (Hakansson and Malcus, 1970; Butler et al., 1991), although computer-aided three-dimensional reconstruction of the duct system in the duck’s salt gland revealed a single anastomosis of the two main ducts at the posterior end of the gland (Marshall et al., 1987). The epithelium of the central canals as well as of the primary and main excretory ducts is composed of two to three cellular layers, with the basal layer

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containing small, rather flat, cells and the superficial layer, large cylindrical ones (Komnick, 1964) (Fig. 6). The latter cells show an abundance of small osmiophilic mitochondria and reveal an enlargement of their lateral surface membranes due to membrane infoldings, and of their apical membrane due to numerous short microvilli (Fig. 6). Marshall and co-workers (1987) differentiated between the central canals lined by cuboidal cells, and the primary and main ducts characterized by columnar cells showing extensive Golgi bodies, lateral membrane interdigitations with apical junctional complexes, and apical microvillar protrusions. Morphometric analysis of the surface area in the duct system, in combination with X-ray microanalysis of intra- and extracellular ion concentrations, led to the hypothesis of the duct system acting as a transporting epithelium (Marshall et al., 1985, 1987). With regard to the soluble enzyme pattern found in the epithelial cells lining the central canals and main ducts in the salt glands of the duck or Herring gull, high activities of G-6-PD, SDH, NADPH-specific ICD, ME, P-hydroxybutyrate dehydrogenase (P-HBDH), and nonspecific acid phosphatase, but not alkaline phosphatase, are expressed. ATPase activity in the epithelium of the duct system proved to be of low activity, thus suggesting no transport function for this glandular compartment (Scothome, 1958, 1960; Ellis et d., 1963; Kuhnel et d., 1969a,b), as hypothesized by Marshall and colleagues (1985). Tubules and collecting ducts of the white fishing sea eagle (Haliaectus leucogaster) and some Falconiform birds are endowed with sulfated and nonsulfated acid mucopolysaccharides (MPS), whose role in secretory or tissue protective processes remains unclear (Sonawane, 1987; More and Sonawane, 1988).

2. Reptilian Salt Glands and Elasmobranch Rectal Gland The duct system of the lacrimal salt gland in the green sea turtle Chelonia mydas consists of central canals that drain the secretory lobes and join to become the main duct. The well-vascularized duct system consists of distal large columnar cells and a proximal pseudostratified epithelium with wide intercellular spaces containing mucocytes (Marshall and Saddlier, 1989). The lateral nasal gland of the desert lizard U . acanthinurus, situated on the ventromedial and lateral aspects of the nasal cavities, drains its secretory fluid via a short excretory duct. It continues as an irregularly branching collecting duct lined by a pseudostratified epithelium whose ultrastructure does not differ significantly from the one described for dark principal cells (Van Lennep and Komnick, 1970). The fine structure of the nonhomologous elasmobranch rectal gland was elucidated by Bulger (1963) and Komnick (1985). Depending on the species, the central duct is lined by a single-layered or pseudostratified epithelium containing a variety of cell types, including mucocytes and interdigitated duct cells, undifferentiated basal and intermediate cells, and granulated cells, possibly of endocrine function (Komnick, 1985). Neighboring cells of the stratified

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epithelium of the rectal gland excretory duct are connected by junctions comprising 12 interconnecting strands, resembling a typical tight epithelium (Claude and Goodenough, 1973).

111. Blood Supply t o the Salt Gland A. Salt Gland Microvasculature Adequate blood supply to the supraorbital salt glands, first mentioned by Jobert (1869) and described in more detail by Marples (1932), represents a necessary prerequisite for active transcellular ion transport. Besides gross morphologic observations and light- or electron-microscope techniques using (ultra)-thin tissue sections (Komnick, 1963b), vascular corrosion castings of the salt gland vasculature, first performed by Fange et al. (1958b) using methacrylate injections into the common carotid arteries and only recently refined (Hossler and Olson, 1990), have proved helpful in unraveling the glandular microvasculature (Fig. 7). Of the marine birds, the salt gland vasculature of only the Herring gull and the saltwater-acclimated Pekin duck has been thoroughly investigated, with some additional information available concerning the nasal salt gland of the Adelie penguin. The salt gland of duck species is supplied by branches of both the ophthalmic and the supraorbital arteries with final drainage of its vascular bed by the ethmoidal and ophthalmic veins (Peaker and Linzell, 1975; Hossler and Olson, 1990). All blood vessels feeding into the glandular parenchyma approach the duck salt gland from its medial surface. Whereas the ethmoidal arterial vessels reach mainly the central and superior portions of the gland, branches of the supraorbital artery supply its temporal portion. With arteries and veins closely intertwined, anastomoses occur between veins and arterioles originating from both the ethmoidal and the supraorbital arteries (Hossler and Olson, 1990). In gull species, the supraorbital salt gland appears to be supplied by the external as well as internal ophthalmic artery (Fange et al., 1958b), with both blood vessels originating from the internal carotid artery. The posterior branch of the internal ophthalmic artery anastomoses with the external ophthalmic artery supplying the caudal portion of the gull’s salt gland, whereas the anterior portion of the internal ophthalmic artery sends off numerous branches to the rostra1 portion of the gland. This arrangement of both arteries inspired Fange and co-workers (l958b) to postulate that “blood could probably bypass the gland via the arterial arch and permit a large reduction in glandular blood flow without reducing the blood flow to the upper beak when the glands are not functioning,” a hypothesis not yet supported by physiologic investigations. In the Adelie penguin, blood vessels probably originating from the supraorbital artery

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pass into the salt gland from the orbit through holes in the frontal bone (Herbert, 1975). Comparing the various anatomical descriptions of avian salt gland blood supply, Hossler and Olson (1 990) presented a scheme whereby the ethmoidal artery was synonymous with the cerebral carotid artery, and the supraorbital artery with the external ophthalmic artery. As mentioned before, the avian salt gland is embedded in a connective tissue capsule with septa diverging into the secretory parenchyma of the gland, thus separating single secretory lobes from each other. Arterial blood vessels proceeding within these interlobular connective tissue bundles merge into arterioles that extend to the connective tissue layers surrounding the central canal (Van Lennep and Young, 1979). A detailed description of these small interlobular arterioles at the electron-microscopic level excluded their participation in the exchange of metabolites between the secretory tissue and the plasma, thus restricting their main task to the sufficient supply of water and electrolytes for the secretory process of the salt gland principal cells (Komnick, 1963b). These arterioles then break up into capillaries winding along single secretory tubules with blood flowing in opposite direction to the flow of the intratubular secretory fluid (Figs. 7 and 8). Each secretory tubule is surrounded by a very dense network of three to seven capillaries (Fig. 2) running toward the periphery of the secretory lobe, equipped with fenestrations 30 to 40 nm in diameter, with short distances between the endothelial cells and the basement membrane of the secretory epithelium (Schmidt-Nielsen, 1960; Ellis et al., 1963; Komnick, 1963b). Near the periphery of the lobes, these capillaries finally pass over into a venous plexus drained by veins in the interlobular connective tissue (Fig. 7). In contrast, the vasculature supplying the epithelium of the central ducts is represented by a simple capillary network. Comparative studies in the salt-secreting gland of a reptile, the chuckwalla Sauromalus obesus, revealed a similar organization of its microvasculature with the stroma surrounding the main excretory duct being penetrated by many large arterioles branching into a dense network of capillaries. These capillaries are closely associated with the principal tubules, with three to six of them surrounding each tubule. The capillaries merge into large venous sinuses situated at the periphery of the gland (Barnitt and Goertemiller, 1985). With regard to the shark rectal gland, microfil injections of the vasculature showed that the single rectal gland artery arising from the dorsal aorta divides into numerous circumferential arterioles. These finally branch into a dense capillary bed running parallel to the secretory tubules to terminate in a large central venous sinus, quite different from the situation in the avian salt gland (Hayslett et al., 1974). Concerning local autoregulatory control mechanisms of avian salt gland microcirculation, fine structural analysis indicated possible local contractility due to the arrangements of collagen and elastic filaments and “passive transformation of endothelial cells” (Komnick, 1963b). Using scanning electron microscopy of vascular corrosion castings, sphincter-like vasoconstrictions have

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FIG. 7 Microvasculature of the avian salt gland. (a) Schematic drawing of the blood supply to a secretory lobe of the duck avian salt gland after ink injection. Arterial blood from an external artery runs in a dense capillary bed from the central ductule toward the rim of the secretory lobe with subsequent venous drainage. (b) Scanning electron micrograph (SEM) of a vascular corrosion cast of a duckling head (dorsal view) showing the supraorbital salt glands (SG).Bar, 200 pm. (c) SEM of a cross section through a salt gland corrosion cast. Large veins (V), interlobular arteries, and secretory ducts (D), as well as several arterioles (arrows) dividing into capillaries are visible. Bar, 200 pm. (d) SEM of the ventromedial surface of a salt gland corrosion cast. Large veins (V),arteries (A), and the venous plexus of single secretory lobes can be distinguished. Bar, 200 pm. ( e ) SEM of

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been observed in a few arterioles close to the point of feeding into the peritubular net of capillaries (Hossler and Olson, 1990).

B. Blood Flow-Secretion Coupling Adaptation of birds kept on freshwater to a hypertonic saline regimen not only resulted in the development of highly specialized secretory tissue, but also coincided with enhanced angiogenesis as shown by morphometric analysis. Thus, angiogenesis after salt acclimation of ducklings is supported by both a marked rise in the weight of glandular corrosion castings and the incorporation of tritiated thymidine into capillary endothelial cells (Hossler, 1982; Hossler and Olson, 1990). In addition, the use of polarographic oxygen electrodes revealed a high arterial blood supply when salt gland tissue had undergone functional hypertrophy due to salt acclimation, and clearly indicated increased arteriolar perfusion with enhanced secretory activity (Fiinge el al., 1963; Peaker and Linzell, 1975). As indicated before, the carotid arteries are almost exclusively responsible for the arteriolar blood supply to the avian salt glands. Recordings of carotid artery blood flow in the Pekin duck using magnetic flow probes yielded values of about 30 ml/min for both blood vessels under thermoneutral conditions (Bech et al., 1982; R. Gerstberger, unpublished observation), indicating that the nonactivated salt glands receive some 5% of the blood perfusing the head region. During systemic osmotic stimulation (0.4 mOsm/min NaCl infusion), carotid blood flow increased to 55 ml/min at an average salt gland-specific perfusion of 15 ml/min for both glands (1.O g tissue weight) (Fig. 8), thus representing 27% of total carotid blood flow due to massive local vasodilation (Gerstberger et al., 1984a; Butler et al., 1989). Stimulated carotid blood flow values were also observed in geese maintained on hypertonic saline as drinking fluid after a systemic injection of 10 ml of a 10% saline or 20% sucrose solution, whereas arterial pressure or heart rate remained unchanged (Burford and Bond, 1968). Graded osmotic stimulation of saltwater-acclimated Pekin ducks ranging from 0.05 to 0.8 mOsm/min induced steady-state salt gland secretion matching the applied salt and water load for all steps chosen. In parallel, arterial blood flow through the glands, as measured using the radioactive microsphere technique, increased with a linear relationship between both functional parameters

the venous plexus of single secretory lobes (corrosion cast) from a duck salt gland. Bar, 200 pm. (0 SEM of a cross section through the gland showing two secretory lobes with the central canals ( m o w ) and the radiating arrangement of capillaries. Bar, 200 pm. (g) SEM of the capillary network supplying the epithelium of the main secretory duct. Bar,SO pm. [(a)From Butler ef al., 1991,copyright 0 1991; ( b g ) from Hossler and Olsen, 1990, copyright 0 1990. All by permission of WileyLiss, a division of John Wiley and Sons Inc.]

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(Kaul et al., 1983) (Fig. 8). Maximal values measured by quantitative radioactive microsphere technique were in the order of 35 ml/min/g wet salt gland tissue, thus representing a more than 20-fold rise in specific blood flow (Kaul et al., 1983; Butler et al., 1989; Gerstberger, 1991). This enormous capacity of the vascular system of the gland could also be verified in freshwater-acclimated geese, where the intravenous application of large volumes of hypertonic saline caused a marked rise in extracellular fluid volume. A subsequent increase in cardiac output at elevated heart rate but unchanged mean arterial pressure was measured. During maximally elevated cardiac output, only salt gland blood flow and coronary circulation were augmented, as determined after 25-30 min. Harderian gland blood flow was not affected by saline loading (Hanwell et al., 1971a,b). For the duck, tight coupling of blood flow and osmolal excretion at a higher time resolution could also be observed when laser-Doppler flowmetry was used to follow local superficial blood flow in the activated salt gland during systemic hypertonic saline administration (Gerstberger et al., 1988; Gerstberger, 1991). This secretion-related rise in glandular blood flow proved to be highly specific for the salt glands, as indicated by unchanged flow values for various vascular beds including brain, Harderian gland, eye, heart, lung, kidney, liver, pancreas, gut, spleen, breast muscle, and web skin (Kaul et al., 1983: Gerstberger, 1991) (Fig, 8). From the proportional increase in blood flow coupled with secretion at a known status of the extracellular fluid compartment, it could be calculated that the secretory epithelium in the duck salt gland removed a nearly constant 20% fraction of the salt delivered to it, producing an arteriovenous difference for "a+] and [Cl-] of 20 and 15 meq/liter, respectively (Kaul et al., 1983) (Fig. 8). The rate of salt gland secretion in the goose appeared to be very loosely correlated with organ blood flow. How exact extraction values for various ions from the plasma (15% for Na+, 21% for C1-, 35% for K+, and 6% for water) could

FIG. 8 Functional aspects of blood flow through the avian salt gland. (a) Schematic drawing of the position of the supraorbital salt gland in the Herring gull. (b) Diagram of the microcirculation in the salt gland of a Herring gull. Countercurrent arrangement of capillary blood flow relative to the flow of secretion within the secretory tubules. (c) Linear relation between osmolal excretion and salt gland blood flow in osmotically stimulated conscious saltwater-acclimated Pekin ducks (SW-AD) (top). and percentage of salt extracted from blood perfusing the salt glands (bottom).(d) Schematic drawing of the unequal distribution of microspheres, injected intracardially, in capillaries of various secretory lobes of the activated salt gland during osmotic stimulation in a SW-AD. (e) Pattern of local salt gland blood flow measured by laser-Doppler flowmetry (flux) during osmotic stimulation in a SW-AD. (0 Organ-specific blood flow (ml/min/g) in various organs of SW-AD during control conditions and systemic hypertonic salt loading (0.4 mOsm/min) as measured by the radioactive microsphere technique. [(a) From Schmidt-Nielsen, 1960. with permission: (b) from Fange et d . , 1958a, with permission: (c) from Kaul ef a)., 1983, with permission.]

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be calculated under these conditions remains to be clarified. The absence of a close correlation of whole organ blood flow and secretion in the goose, as demonstrated for the Pekin duck, might have been due to (a) the manner of OSmotic stimulation not allowing the animal to reach steady-state secretion and (b) the Saphirstein dilution technique, with its inherent inaccuracies such as time delay or tissue preparation (Hanwell et al., 1971b). Despite this generally tight linear relationship between whole organ blood flow and secretion rate in the duck (Kaul et al., 1983), investigations canied out under conditions of high time resolution, employing the recording of local salt gland blood perfusion or the application of various (anti)-secretagogues (see Section VII), revealed partial uncoupling of both blood flow and secretion. Locally measured lobular perfusion varied from almost total vasoconstriction to maximal vasodilation with all intermediate levels possible. The additional observation of vasomotion-like vasoconstrictions and vasodilations during the recording of local “single lobe” salt gland blood flow by laser-Doppler flowmehy, in combination with the uneven distribution of trapped microspheres in various secretory lobes at a fixed time during ongoing secretion, favors the idea of redistribution of intraglandular blood flow to active lobes at a given time with constant whole organ blood perfusion (Gerstberger, 1991) (Fig. 8). Thus the avian salt gland can (a) quickly respond to variable demands by pumping sodium and chloride against a high concentration gradient and (b) alter transcapillary fluid exchange locally due to the vasomotion of precapillary arterioles (Meyer and Intaglietta, 1986). Partial uncoupling of both parameters under various physiologic and pharmacologic conditions is also reported for other exocrine gland systems, with studies in the nonhomologous salt-secreting rectal gland yielding conflicting results. On the one hand, stimulated secretion at increased perfusion rate of an in vitro preparation and diminished excretion of chloride after reduction of the perfusion flow have been reported (Hayslett et al., 1974; Shuttleworth and Thompson, 1986). Also, using an in vivo system, augmented blood flow to the rectal gland during isotonic intravascular volume expansion at unchanged arterial pressure was demonstrated, with secretion being enhanced subsequently (Solomon et al., 1984b, 1985a). On the other hand, treatment with somatostatin or the Na-2Cl-K transport blocking agents bumetanide and furosemide suppressed stimulated secretion in the shark rectal gland, while leaving the glandular blood flow unaffected (Shuttleworth, 1983b; Solomon et al., 1984b). Exocrine secretion independent of arterial blood supply has also recently been described for gastric acid secretion in mammalian species under conditions of high blood perfusion. At low blood flow rates with possibly insufficient oxygen delivery to the stomach mucosa, however, acid secretion is linearly dependent on mucosal blood flow (Perry et d., 1983; Holm-Rutili and Berglindh, 1986; Guth and Leung, 1987). In cat salivary glands, vasoactive substances often cause marked vasodilation after intraarterial injection without concomitant secretion (Lundberg, 1981).

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IV. Salt Gland Innervation A. Nerve Supply t o the Salt Gland 1. Gross Morphology Exocrine glands such as the salivary, lacrimal, Harderian, and bronchial glands, and the pancreas, are known to be innervated by the autonomic nervous system, with the conduction pathway from the central nervous system being made up of preganglionic and ganglionic neurons. Preganglionic fibers emerging from the brain stem in cranial nerves extend through rami of the respective nerves into peripheral ganglionic collections of neurons such as the submandibular, sphenopalatine, or ciliary ganglion, often called parasympathetic ganglia, representing the synaptic sites. With regard to avian salt gland innervation and according to early anatomical observations in the goose, the ramus ophthalmicus of the fifth cranial nerve, before leaving the orbita of the skull, gives off side branches. Two of them enter the orbital ethmoidal ganglion located between the orbital wall and the Harderian gland (Cords, 1904). In addition, a nerve containing fibers of the anterior branch of the nervus facialis, the seventh cranial nerve, and the nervus glossopharyngeus as well as sympathetic fibers enters the ganglionic cell mass. Nerve branches leaving the ganglion diverge to the salt gland, the Harderian gland, or the frontal orbital edge (Cords, 1904). Serial sections of this region in the salt glands of the Pekin duck and the Herring gull did not reveal any connections between the fifth cranial nerve and the secretory nerve (Hakansson and Malcus, 1969; Cottle and Pearce, 1970) (Fig. 9). Despite lack of information due to the inaccessibility of the salt gland innervation, the anterior branch of the seventh cranial nerve must be considered the true secretory nerve, at least in the duck, goose, and gull. Gross morphologic investigations in the Adelie penguin, however, favored the hypothesis of a branch of the fifth cranial nerve innervating the salt glands, with numerous small nerve branches, possibly postganglionic in nature, passing around the frontal edge of the gland (Herbert, 1975). In lizards, the nasal salt glands opening into the anterior chamber of the nasal sac appeared to be innervated by a branch of the lateral ethmoidal nerve or the opthalmicus profundus (Oelrich, 1956; Duvdevani, 1972).

2. Nerve Stimulation-Secretion Coupling Functional indications for the efferent innervation of the salt glands playing a major role in the control of cellular Na+ and C1- transport could be derived from the suppressive action of anesthesia on salt gland blood flow and secretion in osmotically stimulated geese (Hanwell et al., 1971b). In addition, electrical stimulation of the secretory nerve (branch of the seventh cranial nerve) in the

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FIG. 9 Fiber innervation of the avian salt gland. (a) Three-dimensional schematic reconstruction of the secretory nerve (SN filaments), the secretory nerve ganglion (SNG), the ophthalmic branch of the fifth cranial nerve (Vth), and postganglionic fibers innervating the salt gland (SG) of a Pekin duck. (b) Light-microscopic cross section of the secretory nerve ganglion of a Pekin duck salt gland with the secretory nerve (SN) emerging (arrow).Bar, 100 pm.(c) Electron micrograph (EM) of a longitudinal section of a neuronal fiber branch protruding into the secretory parenchyma of a Japanese swan goose salt gland. Bar, 1 pm. (d) Nerve fiber endings on secretory nerve ganglion cells (Bielschowsky staining), revealing button-like varicosities (arrows). Bar, 20 pm. (e) EM of numerous

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anesthetized gull resulted in copious secretion with sodium concentrations as high as 900 meq/liter (Fange et a/., 1958a). To the contrary, the unilateral denervation of one salt gland led to a reduction in blood flow and secretion of the ipsilateral gland compared to the contralateral gland with intact innervation, suggesting that functional parasympathetic innervation is a necessary prerequisite for salt gland function (Hanwell et al., 1971b). The parasympathetic nature of efferent salt gland innervation was deduced from histochemical techniques (see Section IV,B). Using acetylthiocholine as substrate, cholinesterase activity was determined in salt gland tissue sections, with no variation in staining density observed between freshwater-adapted and short-term saltwater-adapted animals (Ash et a/., 1969; Fourman, 1969). Neural control of hypertonic salt secretion against a concentration gradient was also demonstrated for the shark rectal gland. Veratrum alkaloids led to a pronounced chloride secretion from isolated shark rectal glands (Erlij et al., 1981; Stoff et al., 1988). These agents are known to depolarize excitable tissues via increased sodium permeability of their plasma membranes in a way that can be blocked by the sodium channel inhibitor tetrodotoxin. In a series of experiments performed in freshwater-acclimated ducks just recovered from deep anesthesia, the complete denervation of the supraorbital salt glands in the duck including the removal of the secretory nerve ganglion resulted in the expected abolition of saline-induced secretion as well as histochemical cholinesterase staining (Ash et a/., 1969). Sectioning of the secretory nerve central to its ganglionic component, however, surprisingly did not cause glandular unresponsiveness to elevated plasma tonicity. The cholinesterase staining was not markedly reduced with choline acetylase activity lowered to less than 10% of control values. The section of the secretory nerve posterior to the ganglionic region did not appear to contain cholinesterases, indicating that the secretory nerve might be “a mixture of somatic afferent and sympathetic post-ganglionic nerves’’ (Ash et a/., 1969) and that the ganglionic cells themselves might possess osmoreceptive functions. The latter hypothesis appeared to be strengthened by the ineffectiveness of large doses of hexamethonium in influencing salt-induced salt gland secretion. The only electrophysiologic recordings of secretory nerve action potentials available indicated the presence of three fiber populations (Cottle and Pearce, 1970). The inaccessibility of these small fibers might have been the reason for

nerve terminals filled with small translucent (cholinergic) and large electron-dense (peptidergic) vesicles, running parallel at the basis of a secretory salt gland cell (Pekin duck). Bar, 0.3 pm. (f) EM of two nerve terminals in close vicinity to the basal infoldings of a salt gland secretory cell (Japanese swan goose). Bar, 0.3 pm. [(a,b,d) From Cottle and Pearce, 1970, with permission; (c) Unpublished material. courtesy of Prof. Dr. W. Kiihnel; (e) From Hootman and Emst, 1981b, reproduced from J . Cell. B i d . (1981). 91, 781-789; (0 from Kiihnel, 1972, with permission.]

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the negative cholinesterase staining reported by Ash el al. (1969) for the preganglionic nerve fibers. An intact nerve supply to the salt gland appears to be necessary not only for its function, but also for the adaptive hypertrophy during salt adaptation of the birds as demonstrated for geese and ducks with unilateral postganglionic denervation. As mentioned before (see Section KC), both glandular hypertrophy and RNA content or RNA/DNA ratio were enhanced in the intact gland compared to the denervated one (Pittard and Hally, 1973; Hanwell and Peaker, 1973, 1975).

3. The Ethmoidal Ganglion The neuronal “relay station” in avian salt gland innervation, as indicated, is represented by the orbital ethmoidal ganglion (Fig. 9), whose existence in avian species such as the duck or goose was denied in the early anatomical studies by Gaupp (1888). This structure, possibly synonymous with the sphenopalatine ganglion (Webb, 1957), was described to have connections with the nervus trigeminus (fifth cranial nerve), the nervus facialis (seventh cranial nerve), the sympathetic system, and possibly the nervus glossopharyngeus (ninth cranial nerve) (Cords, 1904; Marples, 1932; Webb, 1957). In the salt gland of the Herring gull, parasympathetic nerve fibers of the ramus palatinus of the facial nerve run parallel to the ramus ophthalmicus of the trigeminal nerve. Numerous synapses occur along the common nerve form the ethmoidal ganglion composed of two major ganglionic cell masses located at the common nerve trunk (distal mass) and the junction with fibers branching off to the salt gland and Harderian gland (proximal mass) (Hakansson and Malcus, 1969). For the duck salt gland, serial histological sections enabled exact tracing of postganglionic nerve fibers entering the parenchyma after “travelling around the opthalmic division of the Vth cranial nerve” (Cottle and Pearce, 1970) (Fig. 9). Ganglionic cells, the majority confined to the region where the secretory nerve becomes associated with the fifth cranial nerve, were also found in postganglionic fiber bundles and among fine bundles within the gland. In the Herring gull, the salt gland is evidently provided with postganglionic fibers originating from the distal ganglion portion (Hakansson and Malcus, 1969). Lightmicroscope investigations did not enable the processes and fiber endings to be followed for long distances, whereas EM revealed the presence of synaptic elements on secretory nerve ganglion cells, indicating that they indeed represent the synaptic sites (Cottle and Pearce, 1970) (Fig. 9).

4. Ultrastructure of the Nerve Endings With regard to ultrastructural aspects of salt gland innervation, larger interstitial nerves composed of several axons surrounded by a Schwann’s cell sheath have

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been reported in the duck salt gland. Nonmyelinated fibers leaving the secretory nerve ganglion with subsequent passage into the anterior portion of the gland have been described for the Herring gull salt gland (Fange et al., 1958a). Fawcett (1962) was the first to describe these nonmyelinated fibers running outside the basement membrane, often penetrating the basement membrane to ramify in the gland parenchyma (Fig. 9) with terminal endings in close spatial relationship to secretory cells as well as to collecting duct cells (Kiihnel, 1972; Hootman and Emst, 1981b). Nerve endings were often found in close association with the base of the secretory cells, separated from them by the basal lamina and a layer of connective tissue only (20 nm) (Fig. 9). Terminal axonal swellings have been described in the salt glands of the swan, goose, Pekin duck (Kiihnel, 1972), Eider duck, and Japanese swan goose (W. Kiihnel, personal communications). Pre- and postsynaptic membrane specializations typical of synaptic contacts between adjacent neuronal cells or the neuromuscular junction were not reported. Both in the perivascular space and independent from blood vessels, nerve endings could be located, with their axoplasm containing neurotubules, neurofilaments, mitochondria, and synaptic vesicles of electron-translucent and electron-dense content and varying size (Kiihnel, 1972; Hootman and Emst, 1981b; Lowy and Emst, 1987) (Fig. 9). The smaller, agranular vesicles of 50-nm diameter are characteristic of cholinergic endings, whereas aldehyde fixation in the absence of ferrocyanide revealed that the larger vesicles (100 nm diameter) appeared to have a dense core separated from the surrounding membrane by a thin electron-translucent region typical for peptidergic synaptic vesicles described in other exocrine glands (Lundberg, 1981).

6. Cholinergic Innervation 1. Histochemistry

As indicated before, the avian salt gland receives mainly parasympathetic innervation, as is typical for other exocrine glands. Histochemically, the positive staining for acetylcholinesterase activity was most often taken as a strong indication of cholinergic innervation, despite some difficulties in the interpretation of results obtained due to the presence of this enzyme in some nonneuronal tissues and the possibility that the substrates employed might also be used by butyrylcholinesterase of nonneuronal origin (Peaker and Linzell, 1975). On the one hand, using acetylthiocholine as substrate (Fig. lo), cholinesterase activity was determined in salt gland tissue sections with no variation in staining density observed between freshwater-acclimated and short-term saltwater-acclimated animals (Ash et al., 1969; Fourman, 1969). Ellis and co-workers (1963), on the other hand, reported a different pattern for both groups of animals, with

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FIG. 10 Cholinergic and catecholaminergic innervation of the avian salt gland. (a) Histochemical demonstration of acetylcholinesterase in a salt gland tissue section of a saltwater-acclimated Pekin duck (SW-AD). Bar, 200 pm. (b) Acetylcholinesterase activity near the periphery of salt gland lobes (SW-AD). Bar represents 200 pm. (c) Fluorescent adrenergic nerve fibers in the salt gland following treatment of tissue sections with formaldehyde vapor. Fibers run parallel with secretory tubules with button-like varicosities. Bar, 100 pm. (d) Receptor autoradiogram of intact salt gland sections incubated with [3H]quinuclidinyl benzilate (QNB), a muscarinic receptor antagonist. Silver grain densities are highest over the basal and lateral surfaces of secretory epithelial cells (arrows). Bar, 5 pm. [(a,b) From Ash e r a / . , 1969, with permission; (c) from Peaker and Linzell, 1975, with permission; (d) from Hootman and Ernst, 1982, with permission.]

cholinesterase-positive nerves running through the interlobular, intralobular, and peritubular connective tissue, interlacing fibers adjacent to the secretory tubules but also some endothelial cells of glandular capillaries. Comparative studies in reptilian salt-secreting glands revealed that the compound branched tubular salt-

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secreting glands of sea turtles receive a dual innervation, with one type of subcapsular nerve fibers staining positive for cholinesterase (Abel and Ellis, 1966; Dunson, 1976).

2. Pharmacologic and Physiologic Characterization To elucidate the physiologic significance of parasympathetic innervation for the directed ion transport activity of salt gland secretory cells, the cholinomimetic agent metacholine was administered to isolated salt gland slices, increasing their oxygen consumption by about 50% whereas an increase in the sodium concentration of the medium inhibited cellular metabolism (Borut and SchmidtNielsen, 1963). Treatment of dispersed salt gland cells with metacholine as a muscarinic agonist also resulted in augmented cellular oxygen consumption and enhanced binding of [3H]ouabain, suggestive of stimulated turnover of intrinsic sodium pumps (see Section V,A) (Hootman and Emst, 1981a). Using structurally polarized sheets of primary secretory cell cultures with transmural resistance values in the order of 300 !2 X cm2, the functional (para)sympathetic innervation of the secretory cells themselves could be tested by measuring alterations in short-circuit current (SCC) across the cell layer during drug application (Lowy ef al., 1985a,b) (Figs. 10,12). To mimic parasympathetic innervation, the addition of the cholinergic agonist carbachol to the abluminal side induced an increased SCC in a positive orientation from the luminal to the abluminal side. This transport-stimulating effect could be abolished by the abluminal application of the muscarinic antagonist atropine (Fig. 12). Cell layers grown in the presence of carbachol revealed a marked agonist-induced desensitization, possibly by down-regulation of the muscarinic receptor (Lowy et al., 1985b). To characterize and quantify acetylcholine receptors in the avian salt gland, the tritiated muscarinic receptor antagonist quinuclidinyl benzilate (QNB) was used as radioligand (Hootman and Emst, 1981b, 1982; Hildebrandt and Shuttleworth, 1991b). Binding to an enriched salt gland membrane fraction proved to be saturable and of high affinity with half-maximal binding values (K,) of 35-40 pM regardless of the osmoregulatory status of the animal. Concerning the regulation of receptor numbers per cell or unit of membrane protein during the process of salt acclimation, contradictory results have been obtained by various authors. Hootman and Emst (1981b), working with ducklings adapted to 1% saline for at least a month, reported an up-regulation of cholinergic receptor molecules from 8800 to 14,000 sites per dissociated cell. Hildebrandt and Shuttleworth (1991b), on the other hand, just recently described a down-regulation with values of 532 fmol/mg protein versus 165 fmol/mg protein, using freshly dissociated cells of control ducklings and animals adapted to a 1% saline regimen for only 48 hr. Competitive displacement studies demonstrated high potency of muscarinic antagonists to displace radiolabeled QNB, with lower

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affinities for muscarinic agonists. The ion transport inhibitor furosemide reduced QNB binding to the muscarinic receptor in the duck salt gland, quite different from measurements in other tissues (Hootman and Emst, 1981b). [3H]Quinuclidinyl benzilate binding sites could be localized autoradiographically in principal secretory cells, primarily along the basolateral membrane surface (Hootman and Emst, 1982) (Fig. 10). Whole animal studies yielded the final proof for a functional cholinergic innervation of the avian salt gland. Acetylcholine administered to the carotid arteries perfusing the supraorbital salt glands of conscious saltwater-acclimated ducks or gulls, thus mimicking cholinergic innervation, induced increased 0smolal excretion (Fange et al., 1958a; Gerstberger et al., 1988). Supporting the idea of Bonting and co-workers (1964) that “the cholinergic mechanism controls the gland indirectly through cholinergic vasodilator nerves” is the finding that acetylcholine also markedly enhanced glandular vasodilation with an unchanged general cardiovascular status of the animal (Gerstberger et al., 1988) (Fig. 12). In addition, ongoing steady-state salt gland secretion was totally inhibited by muscarinic antagonists such as atropine or tridihexetylchloride. Glandular blood flow as measured by laser-Doppler flowmetry or the radioactive microspheres technique was reduced to a low atropine-resistant level (Kaul et al., 1983; Gerstberger et al., 1988), as already observed by Fange and colleagues (1958a). The ineffectiveness of atropine to inhibit salt gland blood flow in the goose (Hanwell et al., 1971b), despite abolished secretion, is difficult to discuss. Only two experiments were performed with low doses of atropine. Thus, atropine may not have been able to interfere with vascular cholinergic receptors, while being effective in blocking the muscarinic receptors associated with the secretory principal cells.

C. Vasoactive Intestinal Polypeptide 1. Immunologic Characterization Dense-core vesicles of 100-120 nm in diameter, often containing peptidergic neuromodulators (Lundberg, 1981), were first shown by Kuhnel (1972) for the avian salt gland to be coexistent with small translucent cholinergic vesicles in the same nerve terminal. Analogous to studies in other exocrine glands, such as salivary glands (Bryant et al., 1976), lacrimal glands (Lundberg, 1981), or the pancreas (Sundler et al., 1978), vasoactive intestinal polypeptide (VIP) was found to be a plausible neuromodulator candidate. Immunocytochemistry using VIP-specific antisera indeed revealed the presence of VIP-positive nerve fibers throughout the salt gland parenchyma with terminals ending in the intimate vicinity of both arterioles and the basal membrane of secretory tubular cells (Lowy el al., 1987; Gerstberger, 1988) (Fig. 11). Beaded VIP-positive fiber

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structures could also be localized penetrating into the elastic medial layer of larger arteriolar blood vessels. Although preganglionic fibers did not stain for VIP-like material, ganglion cells and postganglionic fibers showed immunoreactivity, indicative of VIP synthesis in the ethmoidal ganglion. The chromatographic separation of peptidergic extracts with subsequent quantification by radioimmunoassay allowed detailed characterization of endogenous VIP in the salt gland of the saltwater-acclimated Pekin duck, whereas other well-known peptidergic neuromodulators involved in body fluid homeostasis were undetectable (Gerstberger, 1988). Electron microscopically, VIP immunoreactivity could be clearly located within the peptidergic dense-core vesicles in the synaptic nerve endings using either the peroxidase anti-peroxidase (PAP) preembedding or the immunogold postembedding techniques (R. Gerstberger and F. Niirnberger, unpublished observations) (Fig. 11). Thus, VIP-like immunoreactivity was localized in vesicles of synaptic profiles otherwise dominated by translucent cholinergic vesicles. The VIP nerves of the avian salt gland may therefore not be of the “ptype” reported by Baumgarten er al. (1980). Comparable to the findings in the duck salt gland, VIP-like immunoreactive nerve fibers could also be localized in the rectal gland of Squalus acanthias. Thick fibers exhibiting VIP-like immunoreactivity were detectable in its fibromembranous capsule, with thinner branches taking off into the glandular parenchyma and running alongside the epithelial cells surrounding secretory tubules (Chipkin et al., 1988a,b). These fibers were nonmyelinated, as demonstrated by EM. Within the nerve terminals and consistently located along the basal aspects of peritubular cells, the reaction product was confined to densecore vesicles 80-120 nm in diameter. The authors argue that VIP released in the capsular zone, which contains major circumferential branches of the rectal gland artery and plexus of smaller vessels supplying the parenchyma, might affect blood flow to the gland (Chipkin er al., 1988b).

2. Pharmacologic and Physiologic Characterization Comparable to the results obtained with cholinergic agonists, coherent sheets of principal secretory cells responded to VIP treatment with dose-dependently augmented ion transport rates, as indicated by stimulated SCC. Short-circuit current values increased transiently even above the sustained plateau induced by cholinergic agonists (Lowy et al., 1987) (Fig. 12). Vasoactive intestinal polypeptide applied close-arterially to the salt glands of conscious ducks under experimental conditions of threshold secretion (Hammel et al., 1980) caused a marked dose-dependent rise in glandular water and salt elimination from the extracellular space at augmented arterial blood flow through the gland (Gerstberger er al., 1988) (Fig. 12). Representing the morphologic and biochemical correlate for the physiologic findings, high-affinity binding sites specific for radiolabeled VIP

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were found to be concentrated at the basal aspects of the secretory tubules surrounding the central canal of a secretory lobe (Fig. 11). Ligand binding could be inhibited by unlabeled VIP and a peptidergic salt gland extract, but not various VIP analogs tested (Gerstberger, 1988). In the shark rectal gland, veratrum alkaloids induced the release of VIP-like material into the venous effluent of the preparation. The suppression of both veratridine-induced secretion and VIP release by tetrodotoxin or neural blockade suggest that a VIP-like peptide might act as functional neuromodulator in the efferent control of the shark rectal gland (Shuttleworth and Thorndyke, 1984; Stoff el al., 1988).

3. VIP-Acetylcholine Interaction Acetylcholine and VIP could be located in different populations of vesicles in the identical nerve terminal, and both agents enhanced arteriolar blood supply to, and osmolal excretion by, the avian salt gland. Putative interactions of both neurotransmitters and neuromodulators were therefore tested in vifro and in vivo. First, pretreatment of the cultured cells with the muscarinic antagonist atropine did not inhibit the V1P-mediated response (Lowy et al., 1987), nor did muscarinic antagonists suppress VIP-induced vasodilation in the duck salt gland (Gerstberger et al., 1988). Second, the coinfusion of both acetylcholine and VIP did not result in superposition of the vasodilatory actions of each neuromodulator alone, whereas stimulated osmolal excretion appeared to be additive, thus excluding interaction of both agents at the same target. The interplay between cholinergic agonists and VIP in the control of glandular functions proved to be even more complex in other exocrine glands such as the cat salivary glands (Lundberg, 1981). Acetylcholine caused both glandular vasodilation and secretion. Different from the avian salt gland, however, the local intraarterial application of exogenous VIP induced an atropine-resistant vasodilation but no salivary excretion. On the other hand, the infusion of VIP antiserum

FIG. 11 Vasoactive intestinal polypeptide (VIP) as putative neuromodulator in avian salt gland innervation. (a) Light-microscope immunocytochemic demonstration of VIP-positive fibers found throughout the secretory parenchyma of the duck salt gland. Bar, 20 pm. (b) Indirect immunofluorescence for VIP-like material in salt gland tissue sections, with positive labeling of nerve terminal aggregates at the base of secretory tubules (ST). Bar, 10 pm. (c) Electron-microscope demonstration of granular vesicles (G) contained in nerve terminals (NT) close to the basal membrane (BM). immunopositive for VIP-like material using the preembedding Sternberger-PAP technique. Bar, 0.2 pm. (d) Postembedding immunogold staining of granular vesicles (G) in salt gland nerve terminals (NT) containing VIP-like material as neuromodulator. Bar, 0.2 pm. (e) Receptor autoradiogram of an intact salt gland tissue section using radioiodinated ‘2sI-labeled VIP as ligand. Highest receptor densities can be localized at the outer portions of the secretory lobes and associated with main arteries. Bar, 500 pm. [(a,b,e) From Gerstberger. 1988, with permission.]

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FIG. 12 Functional nervous control of avian supraorbital salt gland function. (A) Response pattern of local salt gland blood flow measured by laser-Doppler flowmetq (upper panel) and osmolal excretion (lower panel) to the intracarotid short-term (10 min) application of acetylcholine (ACH), vasoactive intestinal peptide (VIP), and norepinephrine (NOR) in saltwater-acclimated Pekin ducks. (B) Alterations in SCC oriented positive from the luminal to the abluminal side of polarized primary cell cultures of the duck salt gland during bath (luminal or abluminal) application of the muscarinic agonist carbachol (CARB), the phosphodiesterase inhibitor theophylline (THEO), the muscarinic antagonist atropine (ATRO), epinephrine (EPI), the P-adrenergic antagonist propranolol (PROP), the Na-K-ATPase inhibitor ouabain (OUAB), or amphoterecin B (AMPHO) to increase membrane permeability for cations. [(B) From Lowy et al., 1985b. with permission.]

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reduced both vasodilation and salivary secretion. Vasodilatory and secretory responses were augmented during coapplication of acetylcholine and VIP. A possible explanation for this potentiation is presented by (a) a marked increase in immunoreactive VIP in the venous effluent of the gland simultaneously with salivary secretion and vasodilation and (b) the enhancement of muscarinic receptor binding in the presence of VIP (Lundberg ef al., 1982). To make matters even more complicated, muscarinic blockade partially reduced organ-specific vasodilation in the cat submandibular gland only at low parasympathetic stimulation, whereas at high parasympathetic stimulation muscarinic blockade even enhanced glandular vasodilation, possibly due to simultaneously augmented release of VIP. Salivary gland secretion was always totally abolished (Lundberg, 1981). A comparable experimental approach for the avian salt gland might yield interesting results with regard to the co-joint action of two “first messengers” at a common target.

D. Adrenergic Innervation 1. Histochemistry The existence of a sympathetic portion in avian salt gland innervation has already been proposed by early morphologists. Using the Falck-Hillarp technique to demonstrate the presence of monoamines, adrenergic fibers could finally be demonstrated, showing a distribution pattern similar to those stained for cholinesterase in the avian salt gland (Haase and Fourman, 1970) (Fig. 10). Electron microscopy revealed that adrenergic and cholinergic nerve terminals were in close proximity to each other. The synapses of the “bouton de passage” type contained either clear vesicles of 50-60 nm (cholinergic) or granular vesicles measuring 80-100 nm in diameter (monoaminergic). Colocalization of both transmitter substances in the same individual axon terminal could not be established. Fluorescent adrenergic fibers could also be detected in the secretory nerve ganglion following treatment with formaldehyde, with fluorescent nerve terminals surrounding nonfluorescent nerve cell bodies, indicative of sympathetic modulation of postganglionic signal transfer (Peaker and Linzell, 1975). Adrenergic depletion due to reserpine treatment of the animals was found to abolish both catecholamine fluorescence at the light-microscopic level and the presence of granular synaptic vesicles (Haase and Fourman, 1970).

2. Pharmacologic and Physiologic Characterization With regard to a possibly functional sympathetic innervation, the incubation of confluent secretory cell cultures with epinephrine, norepinephrine, or the padrenergic agonist isoproterenol resulted in a dose-dependent stimulation of transepithelial ion transport, with isoproterenol revealing the highest potency.

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The adrenoceptor-mediated response was fully abolished after abluminal application of the P-adrenergic antagonist propranolol, but not the a-specific antagonist phentolamine (Fig. 12). Cultures pretreated with propranolol remained refractory to subsequent incubation with epinephrine, whereas cultures desensitized for muscarinic agonists remained responsive to P-adrenergic stimulation (Lowy et al., 198513; Lowy and Emst, 1987). The authors, however, did not rule out the presence of a-adrenoceptors, which could be demonstrated throughout the salt gland parenchyma employing receptor autoradiography with the radioiodinated a,-agonist iodoclonidine as ligand (Muller, 1991). It would appear that a-adrenergic receptors are present in the avian salt gland and cause vasoconstriction and inhibition of secretion, since epinephrine, norepinephrine, and cervical sympathetic system stimulation reduced salt gland blood flow at increased arterial pressure (Fange et al., 1958a, 1963). In the Herring gull, intravenously infused epinephrine transiently blocked salt gland secretion, whereas phenoxybenzamine facilitated secretion (Fange er al., 1958a; Douglas and Neely, 1969). To simulate possible sympathetic innervation, rather than the unlikely hormonal action of catecholamines upon salt gland function, norepinephrine was administered to the blood perfusing the actively secreting salt glands of saltwater-acclimated Pekin ducks. At increased arterial pressure, steady-state salt gland secretion was only slightly inhibited at strongly reduced arteriolar perfusion of the gland (Gerstberger, 1991) (Fig. 12). A possible synopsis of the functional adrenergic innervation of the avian salt gland might be obtained by taking the simultaneous activation of both inhibitory presynaptic a,-adrenoceptors and stimulatory postsynaptic P-adrenoceptors into consideration. This hypothesis would be in agreement with (a) the marked labeling of salt gland tissue sections by a,-specific ligands (Muller, 1991). (h)the rather low potency of the nonselective a-adrenergic antagonist phenoxybenzamine to displace a,-specific ligands, ( c ) the low physiologic potency of phenoxybenzamine to diminish salt gland blood flow or secretion, but to facilitate ongoing secretion at high drug concentrations (Fange et al., 1958a; Douglas and Neely, 1969; Kaul et al., 1983; Muller, 199I), and (4 the transport-stimulating activity of P-adrenergic agonists in a salt gland tissue culture system (Lowy and Emst, 1987).

V. Secretory Mechanism The precise nature of the cellular mechanisms underlying the elaboration of a hypertonic secretion by salt gland epithelial cells is far from being completely understood, and several models have been advanced by various investigators to explain the extraction of NaCl from blood to lumen, against a large concentration gradient. In addition to the in vivo approach, a variety of in vitro techniques

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have been applied in studying the avian salt gland secretory mechanism, with special emphasis being placed upon the elucidation of intracellular ionic concentrations and fluxes. Although there are significant disparities between the postulated models, one consistent characteristic is that the enzyme Na+- and K+dependent adenosine triphosphatase (Na-K-ATPase) plays an integral role.

A. Na-K-ATPase The salt glands of several avian species have been found to be rich in Na-KATPase (Hokin, 1963; Emst et al., 1967). The enzyme molecule is thought to be a structural component of the plasma membrane, and in all cells its physiologic function is to extrude Na+ from the cell in exchange for K+ (Baker, 1972; Schwartz et al., 1975). Na-K-ATPase can be inhibited or blocked by the cardiac glycoside ouabain not only in avian salt glands but also in other salt-secreting epithelia, including the shark rectal gland (Skou, 1965; Schwartz et al., 1975; Silva et a/., 1977; Kirschner, 1980). The use of this pharmacologic tool has yielded valuable information about the molecular structure, regulation, and cellular distribution of Na-K-ATPase and its function in the salt-secreting process. Procedures for the purification of Na-K-ATPase from the duck salt gland, and also the shark rectal gland, have been determined, and the active form of solubilized salt gland Na-K-ATPase with a molecular mass of 265 kDa appeared to comprise of two a-subunits and two P-subunits with molecular masses of 96-98 kDa and 54 kDa, respectively (Stewart et al., 1976; Lingham et al., 1980; Smith, 1988; Skou and Esmann, 1988) (Fig. 13). Biochemical analysis indicated specific interactions of the protein with certain phospholipids for the Eider duck and Hemng gull (Karlsson et al., 1974). Spin-label studies performed on the Pekin duck salt gland Na-K-ATPase suggested the stoichiometric association of 66 lipids with the intramembranous surface of the Na-K-ATPase molecule (Esmann et al., 1985; Esmann, 1986, 1988; Esmann and Skou, 1988). Ion-dependent protein phosphorylation studies of detergent-purified membranes revealed the marked labeling of the 96- to 98-kDa Na-K-ATPase protein subunit in the presence of Na+, but absence of K+. Highly specific, affinity-purified polyclonal antisera against the Na-K-ATPase a-subunit recognized a 96-kDa protein exclusively (Stewart et al., 1976; Russo et al., 1987) (Fig. 13). In freeze-fracture replicas, avian salt gland Na-K-ATPase was visualized, forming aggregated intramembranous protein particles 8-10 nm in diameter, arranged into clusters and strands. Integration of the solubilized particles into artificial lipid vesicles also resulted in randomly oriented 8- to 10-nm particles (Gassner and Komnick, 1983) (Fig. 13). Binding studies using radiolabeled ouabain in the presence of cyclic AMP and phosphodiesterase inhibitors such as theophylline revealed the existence of at least two Na-K-ATPase subtypes in salt-secreting cells with cyclic AMP-induced transition of site I1 from a negative

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FIG. 13 Sodium-potassium ATPase (Na-K-ATPase) of the secretory cell. (a) Ion-dependent phosphorylation of the 96-kDa Na-K-ATPase subunit (arrowhead) purified from the duck salt gland in the presence of ( I ) Tris buffer alone, (2) Tris buffer + 120 mM NaCI, (3) Tris buffer + 120 mM NaCI + 20 mM KCI, or (4) Tris buffer + 20 mM KCI. Lanes 5-7 are derived from SDS-purified membranes in the presence of ( 5 ) Tris buffer alone, (6) Tris buffer + 120 mM NaCI, or (7) Tns buffer t 20 mM KCI. (b) lmmunoprecipitation of the 96-kDa protein (arrow) in its phosphorylated form ( I ) , and dephosphorylated in the presence of 120 mM NaCl (2) or 20 mM KCI (3). (c) Negative staining of unfixed purified Na-K-ATPase membranes from the duck salt gland. Surface particles are arranged into clusters and strands. Bar, 0.05 pm. (d) Na-K-ATPase-specific histochemical recation product is localized almost exclusively to the intracellular site of the plasma membrane in the basal area of a secretory cell. Bar, 0.3 pm. (e) Alteration in the plasma membrane surface of

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to a noncooperative binding state (Marver et a f . , 1986). When subcellular fractionation procedures and quantification of marker enzymes were employed, the preparation of a membrane fraction from salt glands enriched in Golgi membranes was achieved. This “Golgi fraction” contained high concentrations of one Na-K-ATPase subtype, suggestive of the enzyme being processed in that section of the endoplasmic reticulum (Addis et al., 1987). With regard to the cellular localization, the presence of a potassium- and magnesium-dependent phosphatase component of the Na-K-ATPase sensitive to ouabain was traced histochemically at the cytoplasmic site of the lateral and basal plasma membranes of principal secretory cells in the duck and Herring gull salt gland. Only small amounts of the enzymatic reaction product were found to be associated with the apical cell surface (Abel, 1969; Emst, 1972a,b; Ernst and Mills, 1977; Barmett et al., 1983) (Fig. 13). Using cytochemical lead techniques, adenosine triphosphate phosphorylase activities could also be located within the mitochondria1 matrix (Abel, 1969). Similar results were obtained in binding studies by coupling of a heme peptide bearing peroxidatic activity to the Na-K-ATPase inhibitor ouabain (Mazurkiewicz et al., 1978), or using radioactively labeled ouabain (Emst and Mills, 1977). Reptilian salt-secreting glands, although showing a basic morphology slightly different from that of avian salt glands with columnar cells lining the secretory tubules and mitochondria-rich tuft cells forming the terminal elements, exhibit comparable localization of Na-K-ATPase (Philpott and Templeton, 1964; Crowe et a f . , 1970; Van Lennep and Komnick, 1970; Ellis and Goertemiller, 1974). In the salt-secreting lacrimal gland of the green turtle (C. rnydas), the end product of the cytochemical reaction for Na-K-ATPase was detected primarily along the plications of the plasma membrane bordering the intercellular channels of the secretory epithelium, whereas luminal membranes and cytoplasmic inclusions were rarely observed (Ellis and Goertemiller, 1976). The same holds true for the shark rectal gland (Eveloff et al., 1979; Dubinsky and Monti, 1986) as well as the gills of the killifish (Stirling, 1976). In contrast, polyvalent antibodies raised against the catalytic a-subunit of NaK-ATPase purified from the duck salt gland reacted with the basolateral as well

cells in a duck salt gland (left panel), in the Na-K-ATPase activity, the binding capacity for tritiated ouabain and the number of mitochondria (right panel) under conditions of euhydration ( C ) ,saltwater adaptation (AD) and subsequent deadaptation (DE-AD). (0 Freeze-fracture of unfixed purified Na-K-ATPase membranes with particle-rich (open arrowhead) and particle-poor (solid arrowhead) fracture faces. Bar, 0.2 pm. (g) Freeze-fracture micrograph with a particle-rich fracture face at higher magnification. Bar, 0.1 pm. [(a,b) from Russo et al., 1987. with permission of John Wiley & Sons, Ltd.; (c,f,g) from Gassner and Komnick, 1983, with permission; (d) from Emst, 1972a. with permission; (e) from Merchant et al., 1985 (modified), with permission.]

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as apical membranes of avian principal secretory cells. Within the cells immunolabeling was partially associated with small vesicles or lysosomes, indicating biogenesis or degradative events. Using the immunogold technique, the reaction product was localized on the surface of the microvilli at the apical membrane, independent of the animal’s physiologic state (Russo et al., 1987). Similar observations were reported for the tuft cells of reptilian lacrimal glands with positive labeling of microvilli projecting into the tubular lumen. The rich endowment of the apical tuft cell membranes with Na-K-ATPase is discussed with regard to the rather high concentration of potassium in the secretory fluid of reptilian salt-secreting glands (Ellis and Goertemiller, 1974). The putative functional importance of apical Na-K-ATPase presence in the avian salt gland remains to be elucidated. It might also have been artifactual or due to the staining of cross-reactive proteins. With regard to possible regulatory influences on salt gland Na-K-ATPase activity, the cellular content of Na-K-ATPase increased when freshwateracclimated birds were given hypertonic saline as their only source of drinking fluid (Ernst et al., 1967; Fletcher et al., 1967; Fletcher and Holmes, 1968) (Fig. 13). Long-term saline treatment resulted in a fourfold increase in Na-K-ATPase activity in salt gland tissue accompanied by a comparable fivefold increase in plasma membrane densities as deduced from morphometric methods (Stewart et al., 1976; Merchant et al., 1985; Russo et al., 1987). The report by Holmes and Stewart (1968) of elevated levels in whole cell RNA and ribosomal concentration during salt acclimation of ducklings (see Section II,C) indicated de n o w synthesis of Na-K-ATPase rather than up-regulation of enzymatic turnover. This is supported by the augmented incorporation of radioactivity labeled amino acids into the catalytic subunit of the Na-K-ATPase using slices or an organotypic culture system of duck salt glands under conditions of salt acclimation. In parallel, phospholipid concentrations were significantly stimulated in isolated membranes (Stewart et al., 1976; Lingham et al., 1980; Mazurkiewicz and Barrnett, 1981).

6. Early Models of Salt Secretion Since the earliest studies of avian salt gland function some 30 years ago, a number of models have been put forward to explain the mechanism by which salt gland fluid is secreted. These models have been reviewed in detail by Holmes and Phillips (1983, and only a brief outline of these hypotheses will be given here.

1. Active Na+ Transport Across the Apical Membrane On the basis of their observation that the lumen of actively secreting salt glands was positively charged with respect to the blood, Thesleff and Schmidt-Nielsen

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( 1962) postulated that the secretory mechanism involved active transport of

Na+ by a pump located in the apical membrane. Since this transport could be blocked by ouabain, it was assumed to depend on a Na-K-ATPase system (Thesleff and Schmidt-Nielsen, 1962; Bonting et af.,1964; Van Rossum, 1966) (Fig. 14). Subsequent studies by two independent investigators produced modifications that incorporated Na+ pumps at both the basal and the luminal cell surfaces. In one model (Hokin, 1967) it was postulated that Na+ is concentrated in the secretory cells by means of a ouabain-insensitive process and that a ouabain-sensitive pump, driven by ATP, extrudes Na+ across the luminal membrane. In the other (Peaker, 1971) it was proposed that the concentration gradient for Na+ is established across the luminal membrane by an electrogenic Na+ pump, with entry of Na+ and C1- into the cell across the basal membrane being achieved by an active mechanism involving the exchange of Na+ for H+ and C1- for HC0,- (Peaker and Stockley, 1973, 1974).

2. Active Na+ lkansport into Intercellular Spaces The localization of Na-K-ATPase to the basolateral membranes using cytochemistry and autoradiography with ['Hlouabain (Emst, 1972a,b; Emst and Mills, 1977) together with the fact that intracellular "a+] in salt gland slices was generally shown to be low (Van Rossum, 1966; Peaker, 1971; SchmidtNielsen, 1976) prompted Emst and Mills (1977) to suggest a model for salt gland secretion that omitted an apical pump mechanism. These investigators postulated that the Na-K-ATPase complex was orientated in a way that made possible the transfer of Na+ from the cell to the intercellular space, followed by its paracellular movement via leaky junctions into the lumen of the secretory tubules. This type of shunt pathway maintains a concentration gradient that enables Na+ to enter the secretory cell passively from the blood, taking with it C1-, which then diffuses into the lumen along a favorable electrochemical gradient. The fallability of the idea of the passive role of C1- in salt gland secretion became apparent with the demonstration that furosemide, a C1- transporter antagonist, inhibited salt gland secretion, as did the substitution of C1- by other anions (Gilmore et al., 1977a.b; Hootman and Emst, 1981a; Ernst and Van Rossum, 1982). Accordingly, the above model of the secretory process was subsequently modified to include a secondary active C1- transport coupled with that of Na+.

3. Two-Stage Process A third type of model, advanced by Ellis and co-workers (1977), suggested that salt gland fluid was produced in two stages and required contributions by the two types of secretory tubular cells. The hypothesis proposed that the principal

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FIG. 14 Transepithelial ion transport mechanism in the avian salt gland. (a) Scanning electron micrograph of salt gland secretory cells in primary cell culture forming intercellular tight junctional complexes (arrows). These cultures can be used to study ion transport. Bar, 2 bm. (b) Trace of the voltage potential measured between the blood and the duct of a duck salt gland during secretory nerve stimulation (white bar) in the absence (A) or presence (B) of the Na-K-ATPase inhibitor Gstrophanthin. (c) Schematic presentation of transport processes involved in the excretion of sodium and chloride against an enormous concentration gradient in the avian salt gland. Basolateral location of the ouabain-sensitive Na-K-ATPase, the Na-K-2Cl cotransporter inhibitable by furosemide and a major K+ conductance sensitive to TEA and Ba2+ ions. CI- channels are located at the luminal membrane, and Na+ penetrates into the lumen via paracellular pathways. (d) Transepithelial resistance of salt gland cells for varying periods of time in primary culture after seeding of cultures with dissociated cells. [(a) From Lowy et al., 1985a, with permission; (b) from Thesleff and SchmidtNielsen, 1962, with permission; (d) from Lowy et al., 1985b (modified), with permission.]

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cells do not secrete the hypertonic effluent of the glands; instead, water is passively reabsorbed through the leaky cell junctions from an isoosmotic fluid secreted by the peripheral cells. The driving force of this water movement is the osmotic gradient that exists within the intercellular space due to the presence of Na+ extruded by pumps of the lateral membrane.

4. Post-tubular modification The three types of theoretical models described above all assume that the hyperosmotic fluid that appears at the salt gland nares is elaborated by the cells of the secretory tubules. One study that disputes this has advanced a model depicting salt gland secretion as a modification of the primary hypoosmotic or issosmotic secretion, occurring in the duct system. Marshall and co-workers (1985) used quantitative electron probe microanalysis to measure luminal and intracellular ionic concentrations in fractured, frozen samples of duck salt glands. The results of this analysis showed that the lumen of the secretory tubules contains a secretion that is either hypoosmotic or approximately isoosmotic to blood, certainly not hyperosmotic. Only as the fluid proceeds along the duct system does it become progressively more concentrated, either by the secretion of salt into the lumen or the reabsorption of water. Employing the same technique, Andrews and co-workers (1983), however, obtained results supportive of an active energy-requiring process of the principal secretory cells consistent with active chloride transport as the basis for salt secretion in this tissue.

C. Current Model Using a cell culture system with confluent sheets of isolated duck salt gland cells developing junctional complexes between cells with concomitant increases in transepithelial resistance after 3 days of culturing (see Section IV), Lowy and colleagues (1985a,b) were able to measure changes in short-circuit current and ionic fluxes produced by various agonists and antagonists of salt gland function (Fig. 14). This also allowed differentiation between the apical and the basal sites of drug action. Results of their studies produced a model for NaCl secretion that very much resembles those thought to exist in other secretory tissues, especially the salt-secreting rectal gland of the dogfish (Kirschner, 1980; Smith et al., 1982; Shorofsky et al., 1984; Greger et al., 1986; Lowy et al., 1989). This mechanism of salt secretion is based upon a secondary Cl- transport mechanism. The active process is dependent on ( a ) a ouabain-sensitive Na-KATPase, (b) a furosemide-sensitive Na-Cl symport mechanism, and (c) a barium- and tetraethylammonium (TEA)-sensitive K+ conductance, all situated at the basolateral membrane (Fig. 14). The importance of the Na-K-ATPase could be deduced from experiments where exogenous stimuli such as cholinergic

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agonists were able to increase the turnover rate of plasmalemmal Na-K- ATPase at a constant number of enzyme molecules. Ouabain binding studies revealed the existence of more than 25 million molecules of Na-K-ATPase per dispersed principal secretory cell (Hootman and Emst, 1981a). Ouabain markedly decreased basal and metacholine-induced cellular respiration, and preferentially inhibited active ion transport as measured via altered SCC (Emst and Van Rossum, 1982; Lowy et al., 1989). Evidence for the coupled basolateral carrier-mediated uptake of Na+ and C1- was obtained from experiments demonstrating profound furosemide sensitivity of the salt gland secretory activity, agonist-enhanced oxygen consumption of salt gland slices and dissociated cells (Hootman and Ernst, 1981a; Ernst and Van Rossum, 1982), binding of vitiated ouabain (Hootman and Ernst, 1981a), and active ion transport (Lowy et al., 1985b, 1989). As in other epithelia, this uptake mechanism may also involve K+ cotransport, i.e., a Na-2CIK system, with the added influx of K+ being balanced by back-diffusion through basolateral conductance channels (Silva et al., 1977; Frizzell et al., 1979; Greger and Schlatter, 1984a,b). Using cell-attached and inside-out patch clamp technology. outward-rectifying K+ channels of large conductance could be identified according to their blockade with both external barium and TEA. These K+ channels could be activated in the presence of muscarinic agonists (Richards ef al., 1989). The inward Na+ gradient established by Na-K-ATPase provides the energy for the Na+-coupled uptake of CI- and finally drives the C1- secretion. The accumulated cellular C1- (Andrews et al., 1983) probably leaves the cell across the apical membrane via regulated C1- conductance channels, which have not yet been demonstrated for avian salt glands but are known to be present in the rectal gland of the dogfish (Greger et al., 1985, 1987a,b; Gogelein et al., 1987). This is supported by recent immunolocalization for a cystic fibrosis transmembrane conductance regulator (CFTR) homolog, which is now considered to be a C1- channel, at the apical membrane of the duck salt gland (S. A. Emst, personal communication) and of the shark rectal gland (Marshall, 1991). The K+ diffusion into the blood and the CI- movement into the lumen generate a lumen-negative transepithelial voltage which then secondarily drives Na+ through the paracellular pathway into the tubular lumen. Recent studies employing the Na+ transport channel blocker amiloride ruled out the possible participation of such a channel in the mechanism of salt gland secretion. In addition it was suggested that the paracellular pathway might be similarly permeable to Na+ and K+, as judged from the fixed relationship between plasma and salt gland fluid Na+/K+ ratios under all experimental conditions (Simon and Gray, 1991). The stoichiometry, apparent from Fig. 14, indicates that for each mole of Na+ actively transported by the Na-K pump, 2 mol of NaCl is secreted. This economy of NaCl transport is also present in the rectal gland of the dogfish (Greger et al., 1986) and the thick ascending limb of the loop of Henle in the mammalian kidney. It may be characteristic of CI--transporting epithelia in general.

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VI. Receptive Systems for the Control of Salt Gland Secretion Over the years numerous investigations have been undertaken to elucidate the factors important in the afferent control of salt gland function in birds. Most of this work has involved the application of various osmotic and volume loads given via differing routes. In the majority of cases these studies have been performed in domestic avian species, in particular the duck. Although a picture has emerged from these studies, a number of inconsistencies have also been apparent, particularly with regard to the relative importance of the different regulating stimuli and the sensitivity of the receptive systems in the various avian species examined. However, it should be pointed out that many of the documented experiments were performed using birds that were not saltwater-acclimated and therefore have poor secretory ability (Holmes et al., 1961; Schmidt-Nielsen and Kim, 1964, Peaker et al., 1973). These birds are not appropriate models for examination of the afferent control of salt gland function. In addition, in view of the fact that osmotic status has such an influence on salt gland activity (see below), some of the quantitative characteristics attributed to them should be treated with caution. This is especially true for birds taken from holding pens and subjected to experimental manipulations without first being in a near threshold or steady state of salt gland secretion. Because of the variations in osmotic status known to exist in individual birds (Simon, 1982; Simon and Gray, 1989), statements such as “a two percent increase in plasma NaCl content is required to initiate salt gland secretion” do not carry sufficient unequivocal information. Notwithstanding, it is now clearly evident that ECFT and ECFV can be distinguished as parameters by which the avian salt glands are controlled, and on which salt gland activity feeds back. For the nonhomologous rectal gland of marine elasmobranchs, although increases in ECFV and ECFT have both been shown to stimulate secretion (Burger, 1962; Solomon et al., 1984a,b, 1985b), the effects of changes in tonicity appear to be indirect, acting via the induction of ECFV expansion. Since the plasma of marine elasmobranchs is slightly hypertonic to the seawater environment in which they live, even small elevations in ECFT further enhance the ongoing inward diffusion of water, resulting in ECFV expansion, which drives rectal gland activity. Isovolemic increases in serum osmolality per se, produced by simultaneous hypertonic saline infusion and hemorrhage, do not stimulate rectal gland secretion (Solomon et al., 1985a).

A. Tonicity The salt gland-stimulating effect of increases in ECFT was convincingly demonstrated in many early investigations (Schmidt-Nielsen et al., 1958;

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McFarland, 1964; McFarland et al., 1965; Lanthier and Sandor, 1967; Ash, 1969; Hajjar et al., 1970; Carpenter and Stafford, 1970), which have been thoroughly reviewed by Peaker and Linzell (1975) and Skadhauge (1981). More recent studies have tried to determine the nature and location of the monitoring receptors. Experiments have been designed specifically to evaluate whether the tonicity receptors are located centrally and/or systemically, and whether they are osmosensitive and/or sodium sensitive.

1. Central vs Systemic Tonicity Receptors The idea that increased plasma tonicity activates salt gland secretion via the stimulation of receptors located in the central nervous system (Fiinge et al., 1958a.b; Schmidt-Nielsen, 1960) has been examined by a number of investigators in analogy to the central nervous location of osmosensitive elements involved in the control of water intake, antidiuretic hormone (ADH) release, and renal antidiuresis in mammals and birds (Vemey, 1947; Andersson et al., 1967; Andersson, 1978; Bie, 1980; Simon et al., 1987). Hanwell and co-workers (1972), using geese that had never seen saltwater, found that the intracarotid (ic) administration of hypertonic NaCl did not produce greater salt gland stimulation than the intravenous (iv) route. Moreover, cross-circulation and perfusion studies showed that a raised NaCl concentration in the blood perfusing the head was ineffective in evoking secretion, suggesting that the plasma tonicity must be monitored elsewhere in the body. Since rapid injection of salt into the right atrium initiated salt gland secretion that could be abolished or inhibited by vagotomy or vagal blockade, it was concluded that the tonicity receptors are located in the heart and that the reflex runs in the vagal nerves. In contrast to the above findings, subsequent studies in ducks have demonstrated that brain tonicity receptors do exist. Hammel and colleagues (1983) compared the salt gland responses of saltwater-acclimated ducks to alterations of carotid artery and systemic venous plasma tonicity. Reduction in the osmolality of the blood going to the head, caused by an ic injection of hypotonic glucose, produced a far more pronounced inhibition of ongoing salt gland secretion than did the same dose of glucose given systematically. To further investigate the cephalic contribution to salt gland control, studies utilizing a technique of constant volume split infusion (Simon-Oppermann and Simon, 1982) were undertaken in saltwater-acclimated ducks during steady-state salt gland secretion driven by hyperosmotic NaCl (ic) and isoosmotic glucose (iv) infusions (Gerstberger el al., 1984a) (Fig. 15). Exchange of both infusion lines, with application of the isoosmotic glucose to the carotid arteries and the hypertonic saline to the peripheral circulation, resulted in a fast 13 mOsm/kg decrease in carotid blood tonicity at unchanged peripheral plasma tonicity and markedly reduced osmolal excretion via the salt glands. Additional exchange of both infusates with hypertonic saline administered to the cephalic circulation caused a

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FIG. 15 Central osmosensitivity and integrative hypothalamic control of salt gland function. (a) X-ray roentgenogram of a duck skull (sagittal section), showing the position of a bouble-barrel cannula (C) inserted into the lumen of the third cerebral ventricle (VIII) for local perfusion of the VIlI with artificial cerebrospinal fluid (aCSF). To demonstrate the localization, the ventricular system was filled with an inert contrast medium. Bar, 5 mm. (b) Sagittal section through the hypothalamus of a Pekin duck (Kliiver-Barrera staining). AC, anterior commissure; AP, anterior pituitary; CP, choroid plexus; ME, median eminence; OC, optic chiasma; PC, posterior commissure. Bar, 2mm. (c) Centrally induced inhibition and stimulation of salt gland secretion in saltwater-acclimated Pekin ducks (SW-AD). Simultaneous infusion of hypertonic NaCl (hatched bar) and isotonic o-glucose (stippled bar) into the carotid arteries (ic) and a wing vein (iv), respectively, during constant steady-state excretion (dotted line). Subsequent switching of both infusions at constant total salt and water administration. (d) Inhibition (A) and stimulation (B)of salt gland secretion rate and osmolality of the secreted fluid in SW-AD at steady-state salt gland excretion during intracerebroventricular perfusion with aCSF made slightly hypotonic (A) or hypertonic (B). [(c) From Gerstberger er al.. 1984a (modified), with permission; (d) from Gerstberger el a/.. 1984b (modified). with permission.]

13 mOsm/kg increase in carotid plasma tonicity with parallel 70-170% enhancement in glandular osmolal excretion. The constancy of whole body loading with salt and water strongly suggests the participation of cephalic osmosensitive elements in the perception of these osmotic stimuli. On the other

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hand, the existence of systemic osmoreceptors or volume receptors could be deduced from the reestablishment of steady-state secretion despite the persisting hypotonic or hypertonic infusion into the carotids, resulting in a 9 mOsm/kg increase and 11 mOsm/kg decrease in systemic plasma osmolality at continuous salt and water administration, respectively. To exclude nonosmotic effects of the glucose solution on the sensitivity of the postulated receptive elements or on transport functions of the secretory tissue itself, the isoosmotic D-glucose was replaced by equiosmolal solutions of other nonelectrolytes, all of them producing identical, temporary inhibition of salt gland activity. Considering the differences between these solutes with regard to cellular permeability and metabolic substrate suitability, it seems likely that the inhibition of salt gland secretion induced by them is solely attributable to their common reduction of carotid blood tonicity (Gerstberger et al., 1984a). The indication that tonicity-sensitive structures controlling salt gland activity and accessible via the carotid vascular system lie within the brain has been evaluated by monitoring salt gland responses in the Pekin duck to alterations of hyi pothalamic ECFT. Although a preliminary hypothalamic microinjection study failed to demonstrate any effects of hypertonic NaCl on salt gland activity (Deutsch and Simon, 1980), a subsequent investigation using an intracerebroventricular (icv) microperfusion technique (Fig. 15) in the same species showed that graded icv osmotic stimulations at the diencephalic level produced graded changes in ongoing salt gland secretory activity (Gerstberger et al., 1984b). Hypertonic icv stimulation enhanced salt gland secretion at three times the sensitivity monitored for the inhibitory responses observed during hypotonic icv stimulations (Fig. 15). Comparable results have been reported with regard to the central control of renal function and ADH release in the conscious duck, goat, monkey, and sheep using both ic and icv application of the hypertonic stimuli (Andersson, 1978; McKinley et al., 1978; Swaminathan, 1980; SimonOppermann and Simon, 1982; Gerstberger et al., 1984b). The fact that the effects of icv osmotic stimulations in the duck experiments were obtained under conditions of strong antidiuresis as well as of significant urinary excretion indicates that the alterations in salt gland function were not secondary to changes in body fluid content caused by changes in urinary output. The perfusion technique employed in this study most likely produced stimuli restricted mainly to the rostro-caudal portion of the periventricular tissue adjacent to the third ventricle (Figs. 15,16). Evidence for the important role in osmoresponsiveness which this area of the avian hypothalamus might play could also be derived from central icv stimulations with regard to ADH release and renal water elimination in the duck, and drinking behavior in the pigeon (Gerstberger et al., 1984b; Thornton, 1986; Kanosue et al., 1992). Light-microscope (LM), EM, and scanning electron-microscope (SEM) investigations of the ependymal and subependymal brain parenchyma revealed the presence of numerous neuronal elements penetrating the ependymal layer

AVIAN SALT GLANDS

FIG. 16 Scanning electron micrographs (SEM) of the third cerebral ventricle (VIII)in the duck brain. (a) SEM of a sagittal section through the brain showing the surface of the whole VIII. AC, anterior commissure; AV3V,antenoventral third ventricular region; CP,choroid plexus; ME, median eminence: OC. optic chiasm; PVO, paraventricular organ; RI, infundibular recess. Bar, 200 pm. (b) SEM of a single ependymal cell showing numerous microvilli and cilia (C)as surface membrane protrusions. Bar, 0.5 pm. (c) SEM of single hexagonal cells typical for the AV3V region, ME, or the subfomical organ, lacking a blood-brain barrier, with limited membrane specializations such as cilia (C). Bar, 1 pm. (d) SEM of neuronal dendritic processes (D)originating from liquor-contacting neurons in the PVO.Bar, 0.5 pm.

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of glial origin and protruding into the third ventricular cerebrospinal fluid (CSF) of various avian species including the duck (Vigh, 1971; Vigh-Teichmann et al., 1971; Mikami, 1975; Takei et al., 1978; Korf et al., 1983; Gerstberger et al., 1989). Whereas most of the ventricular lining is composed of specialized ependymal cells with numerous surface cilia and microvilli (Fig. 16), these so-called CSF-contacting neurons occur primarily in areas for which morphologic criteria predict receptive functions, such as the circumventricular organs (CVO) lacking a blood-brain barrier (BBB) or regions like the anteroventral third ventricular zone (AV3V) involved in autonomic control circuits (Vigh and Vigh-Teichmann, 1973; Leonhardt, 1980; Korf et al., 1982; Panzica et al., 1986; McKinley et al., 1991). Scanning electron-microscope studies of the third ventricular wall in the duck and quail revealed a hexagonal shape of cells poor in surface specializations in these regions with bulb- or knoblike dendritic endings sent into the ventricular lumen (Fig. 16). Tanycytes can be traced at the LM and EM level to contact both CSF and subependymal neuronal elements (Korf et al., 1983; Panzica et al., 1986) (Fig. 17). Although all of the afferent projections of these putative receptive neurons have not yet been identified, some of them are known to monosynaptically connect to magnocellular neurons of the paraventricular nucleus (PVN) as demonstrated by retrograde transport of horseradish peroxidase injected into the PVN (Korf et al., 1982) (Fig. 17). These periventricular neurons are therefore thought to be implicated in the release of antidiuretic hormone caused by icv hypertonic stimulation (Gerstberger et al., 1984b; Simon et al., 1992). To demonstrate directly osmosensitivity as a property of neurons located particularly in the periventricular layer, being labeled by retrogradely transported tracers applied to the PVN, single unit recordings were performed. These extracellular in vitro recordings were obtained from neurons in various hypothalamic areas of the duck brain using a duck hypothalamic slice preparation (Korf et al., 1982; Kanosue et al., 1990). Accordingly, hypertonic stimulation due to slightly elevated NaCl concentration in the artificial CSF used for brain slice incubation excited 25 and inhibited only 4 of 48 periventricular layer neurons. Hypotonic stimulation inhibited 9 of 10 neurons activated by hypertonic stimulation (Fig. 17). Osmoresponsive units in other areas of the hypothalamus such as the magno- or parvocellular PVN were rarely observed in the duck. In the rat brain, however, intrinsic osmoresponsiveness of hypothalamic cells could be demonstrated for the supraoptic nucleus (SON), organum vasculosum laminae terminalis (OVLT), or AV3V region, with the formation of an osmoreceptive complex (Honda et al., 1990). Afferent connections of this periventricular osmosensitive zone to salt gland-regulating centers cannot be excluded; on the contrary, they are very likely. Moreover, the additional, yet unidentified, location of salt gland tonicity receptors in brain structures that are accessible to both CSF and blood, e.g., CVO structures, would be compatible with the experimental data.

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FIG. 17 CSF-contacting neurons as putative central osmosensors. (a) Small neurons in the periventricular layer (PL) of the third cerebral ventricle (VIII) are retrogradely labeled by horseradish peroxidase microinjected into the magnocellular portion (MC) of the paraventricular nucleus. Bar, 100 pm. (b) Golgi impregnation of a frontal section through the duck hypothalamus showing a magnocellular neuron (MCN) of the paraventricular nucleus with its dendrites connected to a neurite (arrows) originating from a subependymal liquor-contacting neuron (CN) close the VIII. Bar, 50 pm. (c) Higher magnification of the subependymal CN. Bar, 20 pm. (d) Osmosensitivity of a neuron located in the periventricular subdivision of the hypothalamic paraventricular nucleus (PL) adjacent to the third cerebral ventricle in the duck brain. Extracellular recordings of neuronal discharge rate in a hypothalamic slice preparation during hypotonic (left) and hypertonic (right) stimulation. [(a)From Korf er al.. 1982, with permission; (b,c) from Korf er al., 1983, with permission; (d) from Kanosue er al., 1990 (modified), with permission.]

2. Osmosensitivity vs Sodium Sensitivity The first report on salt gland function reported that iv infusion of sucrose stimulated a cormorant to produce a nasal gland secretion of a concentration similar to that induced by an infusion of NaCl solution (Schmidt-Nielsen et al., 1958). The indication of this finding that any elevation of the osmotic concentration of the blood will stimulate salt gland secretion appears to have been subsequently confirmed by the observation that an increase in the plasma concentration of compounds as diverse as mannitol, LiCl, KCl, and arginine-HC1 can activate

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avian salt glands (Hajjar et al., 1970; Hanwell et al., 1972; Deutsch et al., 1979; Erbe et al., 1988). However, salt glands do not respond to hyperosmotic doses of MgCl, or urea (Carpenter and Stafford, 1970; Deutsch et al., 1979); therefore, a simple increase in the osmotic concentration of the plasma does not unequivocally drive salt gland activity. As originally proposed by Vemey (1947), a tonicity receptor is thought to be stimulated by a reduction in its cell volume, caused by an efflux of water down an osmotic gradient created by a rise in extracellular solute concentration (Hammel, 1981). Thus, if all other factors are equal, the receptor-stimulating potency of a particular blood solute should be inversely proportional to the rate at which it enters the receptor. For systemic receptors, solute specificity would be determined solely by its receptor membrane permeability. For central receptors, solute permeability of the BBB also needs to be taken into consideration. With regard to the putative systemic tonicity receptors, available information indicates that they are generally osmosensitive. Accordingly, they can be stimulated by hypertonic mannitol and sucrose, which are by and large excluded from receptive cells, but not significantly affected by hypertonic urea, which permeates cell membranes. As for central receptors, the situation is somewhat more complex, with interpretation depending on whether they are thought to be situated on the blood (e.g., CVO) or the brain side of the BBB. However, available data can be taken to indicate that receptors are present in both locations, and that they are predominantly Na+ (or cation) sensitive. Accordingly, salt gland activity driven by the ic infusion of hypertonic NaCl (3000 mOsm/kg) is markedly decreased when the saline is replaced by equiosmolal sucrose (Gerstberger et al., 1984a). The reduction of the "a+] in the cephalic compartment, at a constantly elevated blood osmolality, appears to be the reason for this response and is compatible with the idea that the relevant Na+ receptors are located on the blood side of the BBB. These or other Na+ receptors involved in salt gland control are also accessible to CSF, since a reduction in CSF "a+] by the icv application of hypertonic mannitol (Deutsch and Simon, 1980) or artificial isoosmotic CSF in which NaCl was replaced by sucrose (Gerstberger et al., 1984b) caused a reduction in ongoing secretion. Recently, evidence has been obtained extending the concept of a specific Na+sensitive receptive mechanism, to include general cations. Osmoresponsive neurons present in hypothalamic slice preparations have been shown to respond equally to stimulation by artificial CSF made hypertonic by either NaCl or LiCl, but not sucrose (Kanosue et al., 1990). Taken together with the fact that the brain receptors thought to be involved in the osmotic stimulation of ADH release in ducks are sensitive to Li+, Ca2+and Mg2+,as well as to Na+, but again not to sucrose or mannitol, the concept of nonspecific cation channels gains support (Deutsch and Simon, 1980; Gerstberger et al., 1984b; Kanosue et al., 1992).

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6. Volume Early investigations concerning the significance of volume influences on salt gland secretion produced no consensus of opinion on the matter (Holmes, 1965, 1972; Hanwell et al., 1972; Ruch and Hughes, 1975; Zucker et al., 1977; Peaker, 1978). More recently, however, clear evidence for the role of a volume component in avian salt gland function has emerged, although some degree of species variation is apparent with regard to sensitivity, particularly between ducks (the most studied avian species) and gulls (Hughes, 1987, 1989). For the analog structure of the shark rectal gland, isotonic ECFV expansion was shown to stimulate CI - secretion using an in situ preparation as well as intact anesthetized animals (Erlij et al., 1980; Solomon et al., 1984a; Erlij and Rubio, 1986). There are a number of examples in which changes in avian salt gland activity have been reported in response to alterations in ECFV, with current emphasis being placed on the relative importance of vascular and interstitial fluid volume (ISFV).

1. Vascular Volume A number of studies examining the effect of hemorrhage on salt gland function have been carried out. In ducks, removal of blood in amounts ranging from 7 to 30% total blood, before (Phillips and Harvey, 1980) or during (Deutsch et al., 1979; Hammel et al., 1980; Simon-Oppermann et al., 1984) salt gland secretion, reduced salt gland activity (Fig. 18). Reinfusion of the removed blood increased ongoing secretion. In gulls, neither the removal of 20% blood volume nor the infusion of 20% additional blood had any effect on salt gland secretion (Hughes, 1987). It has been suggested, at least in the case of ducks, that an increase in body fluid content might diminish the threshold tonicity for salt gland activation (Hammel et al., 1980; Simon, 1982). Consequently, vascular expansion by iv infusion of isoosmotic saline with or without dextran or slightly hypotonic NaCl induced salt gland secretion (Zucker et al., 1977; Hammel et al., 1980; Gray et al., 1986). Evidence for this volume effect to be acting via a change in the secretory threshold tonicity originated primarily from experiments in which the composition and volume of the body fluid in Pekin ducks were altered by iv infusions of various NaCl solutions (Hammel et al., 1980) (Fig. 18). Control experiments consisting of an iv infusion of hypertonic saline induced rates of secretion of salt and water by the salt glands that matched the rates of infusion, when the ECF "a+] exceeded some preestablished threshold concentration ("a+],,). Hydration of the ducks with an infusion of hypoosmotic saline, with the same amount of salt applied as during the hypertonic stimulation, had the effect of reducing ECF "a+] by 8 meq/liter and expanding ECFV due to the retention of 72% of the water infused. Repeating the control infusion of hyper-

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FIG. 18 Afferent volume regulatory and hypothalamic integrative control of salt gland function. (A) Inhibitory and stimulatory actions of isotonic extracellular volume alterations (bleeding, reinfusion of blood) on salt gland osmolal excretion in the saltwater-acclimated Pekin duck (SW-AD) during steady-state secretion (dotted line). (B) Nasal salt gland excretion in SW-AD (solid line) in response to hyperosmotic (0.4 ml/min of 1000 mOsm/kg NaC1) (A,C) and almost isotonic (1.4 ml/mm of 277 mOsm/kg NaCI) (B) infusions into the systemic circulation (dotted line). (C) Effect of graded hypothalamic cooling on salt gland activity in a SW-AD receiving a continuous systemic hypertonic salt load. (D) Effects of hypothalamic cooling and warming on salt gland activity in an Adelie penguin receiving a continuous systemic hypertonic salt load (dashed line). [(A) From Simon-Oppermann et al., 1984 (modified) with permission; (B) from Hammel et al., 1980, with permission; (C) from Simon-Oppermann et al., with permission; (D) modified from “The Proceedings of the Third SCAR Symposium on Artic Biology” by H. T. Hammel, J. E. Maggert, E. Simon, L. Crashaw, and R. Kaul. Copyright 0 1977 by Gulf Publishing Company, Houston, TX. Used with permission. All rights reserved.

tonic saline under these conditions indicated no significant change in the ratio of the amount infused and secreted. Since ECF “a+] had been reduced by the hypoosmotic infusion, the conclusion reached was that the increase in ECFV lowered the “a+],, for secretion. Further evidence for the putative inverse relationship between “a+], and ECFV could be obtained by comparing the ratio of the salt secreted to that infused before and after removing 15% of the total blood, showing that ECFV depletion reduced salt gland secretion by 24%.This result is in good agreement with results on ducks (Deutsch et al., 1979; Kaul

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and Hammel, 1979), which demonstrated that dehydration-induced ECFV reduction raised the “a+],, for salt gland secretion.

2. Interstitial Fluid Volume From the above experiments it is impossible to evaluate which, if any, compartment of the ECF, i.e., vascular or ISFV, is the most important in salt gland regulation, since the induced changes occurred in both. To make possible the evaluation of the contribution of both ECF compartments in afferent salt gland control, the salt gland response to iv dextran was monitored. The salt glands showed a diminished secretory response to a hyperosmotic (Hammel et al., 1980) and isoosmotic (Keil, 1990; Keil et al., 1991) NaCl load administered together with hyperoncotic dextran solutions. The substantial reductions in hematocrit values after dextran infusion clearly indicated a considerable rise in the intravascular volume, at the expense of the extravascular volume. These findings have been interpreted to indicate that salt gland activity is more dependent on the ISFV than the vascular volume (Simon, 1982; Keil et al., 1991). However, although the resultant salt gland inhibition conforms to the view that the activity of these glands is influenced by the ISFV, the fact that secretion continued (at a reduced level) with the infusion of an isotonic volume load (Keil et al., 199I), which is presumably driven by intravascular expansion, makes it impossible to determine which compartment is more important. Rather it is more accurate to say that salt gland activation by the ECFV is dependent on the algebraic sum of excitatory and/or inhibitory stimuli arising from both the intravascular and the extravascular compartments.

3. Extracellular Fluid Volume Receptors The implication that changes in the ECFV affect salt gland function raised the questions where and how these changes are monitored and transduced. Volume is an extensive property and since only intensive properties of the body may be transduced into neural signals, tension receptors somewhere in the ECF must monitor the size of that space and elicit the salt gland effects produced by changing the ECFV. In the case of intravascular volume, stretch receptors possibly located in the vicinity of the heart with afferent fiber passage in the sensory component of the nervus vagus (tenth cranial nerve) connecting the transducers with higher brain stem centers might be postulated. Vagal transsection or anesthetic blockade markedly inhibited ongoing salt gland secretion (Hanwell er al., 1972; Gilmore et al., 1977b; Simon-Oppermann et al., 1980). Concerning the extravascular volume, nothing is known about either the structure or the location of the postulated interstitial tension receptors. With regard to the afferent control of volume-induced rectal gland function in the dogfish or salt gland activity in various reptiles, only limited data are

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available about localization and characterization of putative volume-sensitive elements involved in the signal perception and transduction processes (Burger, 1962; Dunson, 1976). The stimulatory effect of localized application of veratridine into the atria or the conus arteriosus of the dogfish heart led to the questionable postulate of a receptive mechanism situated in the vicinity of the heart to monitor changes in ECFV or vascular filling in these animals (Erlij and Rubio, 1986). Preliminary studies performed in various sea turtle species of the genus Chelania indicate that secretion of the lacrimal salt glands is not initiated and controlled by increased ECFV (Hudson and Lutz, 1986; Marshall and Cooper, 1987; Nicolson and Lutz, 1989). Recently Hammel (1989) raised the interesting possibility that the neural control of avian salt gland secretion is enhanced and sustained by autofacilitation. On the basis of the fact that soon after a salt gland response has been initiated by an iv infusion of salt the rate of excretion exceeded the rate of infusion, Hammel argued that salt gland activity must be sustained by some unknown facilitation of the neurons regulating secretion, even as the initiating stimuli return to threshold. Further evidence for autofacilitation by a positive feedback loop was indicated by the finding that once the activity of the regulating neurons is sustained by their own activity, they are less easily inhibited by regulatory hormones or ISFV reduction due to iv dextran administration. However, when inhibitory hormones or dextran accompany a saline infusion driving salt gland secretion from the onset, autofacilitation is impaired.

C. Neural Integration The multiplicity of afferent inputs regulating salt gland function (cerebral and extracerebral osmoreceptors, body fluid volume receptors) indicates the need for an integrating center in the efferent control. Most of the available evidence indicates that this is accomplished by hypothalamic neuronal networks, which in addition may also integrate salt gland activity with other regulatory systems including those of kidney function, cardiovascular control, and thermoregulation. Afferent control of salt gland secretion is known to be closely coordinated with that of kidney excretion (Skadhauge, 1981; Simon, 1982; Simon-Oppermann and Gerstberger, 1989; Simon and Gray, 1991), with the receptive elements controlling both systems probably being colocalized but not identical (Gerstberger et al., 1984b). The presumably osmosensitive afferents from the wall of the third ventricle, as well as ascending information from the vagal and glossopharyngeal nerves, were shown to converge in the PVN at sites where microelectrostimulation elicited antidiuretic reactions. Vagal and glossopharyngeal afferents are relayed in sensory nuclei of the avian brainstem such as the nucleus tractus solitarii (NTS) (Korf et al., 1982; Korf, 1984). Neurons from this region of the hypothalamus apparently descend to the parasympathetic brain-

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stem nuclei efferently innervating the salt glands. This can be inferred from the observation that hypothalamic cooling inhibited ongoing salt gland activity in penguins and ducks (Hammel et al., 1977; Simon-Oppermann et al., 1979), whereas hypothalamic w m i n g transiently activated penguin or duck salt glands (Hori et al., 1986) (Fig. 18). The existence of a temperature dependence of signal transmission in hypothalamic neural integration supports an additional integration between osmo- and thermoregulation at this level. With regard to possible agents involved in the hypothalamic integration of various afferent osmoregulatory and volume-regulatory signals, there are clear indications that central neuromodulators such as norepinephrine or angiotensin I1 (ANGII) play a major role in the homeostatic control of salt gland secretion. The avian hypothalamus is strongly innervated with tyrosine hydroxylase- and dopamine P-hydroxylase-positive fibers, two key enzymes in the synthesis of brain catecholamines. The existence of 01- and P-adrenergic receptors in hypothalamic structures involved in central osmoperception and integration, such as the PVN, the anterior hypothalamus, or the periventricular zone, is consistent with the postulated hypothesis (Simon-Oppermann and Giinther, 1990; Gerstberger et al., 1992; Miiller and Gerstberger, 1992) (Fig. 19). A brain-intrinsic ANGII system has been detected in the saltwater-acclimated Pekin duck, with ANGII being regulated in a clearly ECFV-dependent manner (Gray and Simon, 1987; Ramieri, 1988). Receptive systems for the peptide could be localized in areas within the BBB such as the PVN, SON, AV3V region or medullary centers such as the NTS or olivary complex, all involved in homeostatic control circuits of salt and water balance or the cardiovascular system (Gerstberger et al., 1987a,b, 1992; Simon et al., 1987, 1992). Although physiologic data concerning the action of brain-intrinsic norepinephrine on avian salt gland function are still lacking, the central application of ANGII is well known to inhibit ongoing salt gland secretion in the duck. This is probably mediated via altered parasympathetic outflow to the glandular vasculature and secretory parenchyma (Gerstberger et al., 1984~).

VII. Hormonal Control of Salt Glands Hormones defined in the classical way as mediators “secreted by specialized endocrine cells into the circulating blood and travelling relatively long distances to targets they act on” (Norris, 1985) have also been extensively investigated with regard to their putative modulatory role of avian salt gland function (Holmes and Phillips, 1985; Butler, 1984; Butler et al., 1989). Emphasis was placed on the pituitary-adrenal axis and various peptide hormones of osmoregulatory importance, such as the antidiuretic hormone AVT, blood-borne ANGII, or the recently discovered atrial natriuretic factor (ANF).

FIG. 19 Receptor autoradiographical demonstration of high-affinity binding sites for neurotransmitters involved in the hypothalamic integration of afferent signals important for the control of avian salt gland function. (a) Distribution of hypothalamic angiotensin I1 binding sites labeled with the radioiodinated receptor antagonist '251-labeled 'sar-*ile-angiotensin 11. (b) Distribution of hypothalamic a,-adrenergic binding sites labeled with the radioiodinated receptor antagonist 251-labeled HEAT (c) Distribution of hypothalamic a,-adrenergic binding sites labeled with the radioiodinated receptor agonist 1251-labeledclonidine. AC, anterior commissure; AM, amygdala complex; AV,V, anterioventral third ventricular region; LS, lateral septum; PVN, paraventricular nucleus; SFO, subfornicnl organ. Bar,2 mm. [(a) From Gerstberger et al. 1987a. with permission; (b,c) Unpublished material, courtesy of Mr. A. Muller.]

,

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A. Pituitary-Adrenal Axis The idea that components of the pituitary-adrenal axis may be involved in the regulation of avian salt gland function originated from the observations that adenohypophysectomy (Wright et al., 1966; Holmes et al., 1972) or adrenalectomy (Phillips et al., 1961; Thomas and Phillips, 1975a,b) inhibited or blocked salt gland responses to hypertonic saline loading. Moreover, birds with the best developed salt glands have the largest adrenals (Holmes et al., 1961; More and Patil, 1988). Salt loading, both acute and chronic, elevated plasma corticosteroid concentrations, although not in every case (Donaldson and Holmes, 1965; Macchi et al., 1967; Allen et al., 1975; Harvey and Phillips, 1980, 1982; Klingbeil, 1985a,b). Corticosterone, as the main naturally occurring adrenocorticoid in avian species (DeRoos, 1961; Sandor et al., 1963; Donaldson et al., 1965), restored salt gland secretion in adrenalectomized birds and augmented secretion in normal animals (Holmes et al., 1961; Holmes, 1972; Harvey and Phillips, 1982). In addition, active salt glands accumulated much greater amounts of corticosterone than other tissues (Bellamy and Phillips, 1966; Takemoto et al., 1975), and two types of corticosterone receptors have been isolated from cytosolic fractions (Sandor and Fazekas, 1973, 1974; Allen et al., 1975; Sandor et al., 1977). Although the above findings seem to indicate a regulatory role for corticosterone in salt gland function, recent observations have produced reasons to doubt that this role involves a direct effect on the initiation or maintenance of salt gland secretion (Butler, 1980, 1984, 1987; Butler et al., 1989). It was shown that adrenalectomized ducks given 0.9% saline as drinking fluid to counterbalance postoperative loss of salt and water, or force-fed to overcome anorexia, had salt glands that functioned almost completely normal within 7 days of adrenal gland removal (Miller and Riddle, 1943; Thomas and Phillips, 1975a; Butler and Wilson, 1985; Butler, 1987; Butler et al., 1989) (Fig. 21). Similarly, the absence of adrenal glands does not block the adaptive hypertrophy associated with transfemng ducks from fresh water to saline (Butler, 1984). Other experimental observations which indicate that corticosterone has no direct action on salt gland activity include its lack of effect in vitro on salt gland slices and the absence in vivo of any alteration in the salt gland secretion of birds given corticosterone acutely or long term (Wilson and Butler, 1980; Butler, 1984; Harvey et al., 1985). The possibility that the effect of adrenalectomy upon salt gland function is related to cardiovascular alterations induced by steroid depletion was evaluated by Butler and Wilson (1985). Although removing the adrenal glands produced no changes in blood volume, there was a gradual reduction of both systolic and diastolic pressures during the first 3 days after adrenalectomy. This decrease was prevented by betamethasone, demonstrating that steroids have an essential role in avian blood pressure maintenance. Accordingly, it appeared that the

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reduction in plasma corticosterone levels caused by hypophysectomy or adrenalectomy produced cardiovascular failure and resultant blood flow insufficiency, which prevented an increase in blood supply essential for salt gland activity (Hanwell and Peaker, 1973; Kaul et al., 1983; Butler et al., 1989). Corticosterone may also have another indirect stimulatory effect on avian salt glands by elevating plasma glucose concentration, which itself has been shown to enhance salt gland secretion by increasing the availability of metabolic substrate for the sodium-transporting mechanism (Peaker et al., 1971; Holmes, 1972; Allen et al., 1975). In addition, it has been suggested that corticosterone is physiologically relevant for the homeostatic adaptations of salt glands by regulating transcriptional events leading to protein induction required by hypertrophying glands (Sandor and Mehdi, 1981; Harvey and Phillips, 1982; Sandor et al., 1983; Harvey et al., 1985; Holmes and Phillips, 1985). The observation that adrenalectomized ducks, without corticosterone, progressively secreted fluid at a higher rate and electrolyte concentration, however, indicates that any role of corticosterone in developing glands is not essential (Butler, 1980). The fact that adenohypophysectomy completely inhibited salt gland secretion, which could be restored by adrenocorticotrophic hormone (ACTH) supplement, is likely to be dependent on the role of ACTH in maintaining plasma corticosterone concentrations rather than any direct effect it may have on salt glands (Wright et af.,1966; Bradley and Holmes, 1971; Holmes et af.,1972). Harvey and co-workers (1985) found that ACTH injection had no effect on salt gland function in ducks, and although in another investigation ACTH did stimulate salt gland secretion, its effect was thought to be due to an elevation of plasma glucose and potassium, both of which can stimulate salt glands (Peaker et al., 1971; Holmes, 1972). In the same study, however, ACTH did reduce nasal fluid “a+] and [K+], an effect not seen with hyperglycemia or hyperkalemia. It may represent a direct influence of ACTH on water movements in the salt gland. For aldosterone, there is no evidence that a direct effect on salt glands is of physiologic significance. Although this steroid has been shown to delay salt gland secretion and reduce the sodium content of the fluid, the duration of secretion is prolonged. Thus, total osmotic excretion is unaffected (Gill and Burford, 1968). Other studies have failed to demonstrate any type of effect of aldosterone on salt gland activity (Phillips and Bellamy, 1962; Holmes, 1972), and it is generally believed that this mineralocorticoid plays no role in salt gland regulation (Thomas and Phillips, 1975a; Harvey and Phillips, 1982; Simon and Gray, 1989).

B. Angiotensin II The octapeptide ANGII, the foremost product of the renin-angiotensin system, effects avian salt gland function in a way that is consistent with its generally

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accepted role of salt and fluid conservation (Peach, 1977), namely inhibition of salt gland secretion. This effect, first demonstrated in ducks by Hammel and Maggert (1983) has been repeatedly confirmed in this species (Butler, 1984; Wilson et al., 1985; Gray et al., 1986) as well as in gulls (Gray and Erasmus, 1989a) (Fig. 20). In each of these studies it was shown that elevation of the circulating level of ANGII reduces the salt and fluid output of active salt glands. In addition, reduction of the systemic concentration of ANGII by blocking its production using captopril, a converting enzyme inhibitor, enhanced extrarenal excretion in saline-loaded ducks (Wilson et al., 1985), although this effect may have involved a reflex compensation for the reduced renal output induced by captopril. The threshold plasma concentration of ANGII for its inhibitory action seems to be about 150 pg/ml (Gray and Erasmus, 1989a), although this is likely to be very much dependent on the drive to stimulate salt gland secretion (Hammel, 1989). This concentration of ANGII is well within the range measured in the blood from various bird species with salt glands (Gray and Simon, 1985; Gray and Erasmus, 1988); therefore the effect of ANGII is clearly physiologic. The inhibitory action of peripheral ANGII on salt gland secretion could be abolished by coapplication of the specific ANGII receptor antagonist 1sar-8ile-ANGII.but not 'sar-8ala-ANGII commonly applied to mammalian species (Hammel and Maggert, 1983). Specific blood flow measurements using the radioactive microspheres technique or laser-Doppler flowmetry in conscious animals clearly demonstrated that ANGII markedly inhibited both salt gland secretion and arterial blood perfusion of the gland at almost unchanged arterial pressure, heart rate, and cardiac output (Butler, 1984; Gerstberger, 1991). The seemingly obvious direct action of ANGII on both the vasculature and the secretory parenchyma of the avian salt gland, however, can be ruled out due to (a) the higher efficacy of intracarotid ANGII application in inhibiting ongoing secretion compared to that of systemic peptide administration, (b) the lack of ANGII-specific binding sites in the salt gland tissue (Gerstberger et al., 1987a,b; Gerstberger, 1992), and (c) the abolition of ANGII-induced salt gland inhibition after ganglionic blockade (Butler et al., 1989). Figure 20 shows that duck salt glands stimulated to secrete by an intravenous continuous infusion of hypertonic saline were switched off transiently by a single injection of ANGII. When steady-state salt gland secretion had been reestablished, blockade of ganglionic transmission with subsequent total reduction of secretory activity could be reversed by iv infusion of metacholine, a cholinomimetic drug. With the glands secreting in the absence of any neural input, ANGII then failed to have any inhibitory effect on their fluid or salt excretion. Blood-borne ANGII did not directly regulate NaCl transport by secretory cells in salt glands, nor did it cause direct vasoconstriction of blood vessels supplying them. The inhibitory action of ANGII was mediated via modulation of the neural control to the salt gland, presumably within the central nervous system. Whether this is limited to a

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reduction of the cholinergic input or also involves a stimulation of sympathetic fibers innervating the glands is unclear. Neuronal entities without a well-established BBB, such as CVOs, represent the prime targets for circulating ANGII to inhibit salt gland function through its central action. Receptor binding studies revealed the presence of high-affinity ANGII binding sites in the SFO and the median eminence of the duck hypothalamus being accessible to blood-borne ANGII, comparable to data available for mammalian species with regard to centrally induced drinking (Gerstberger et al., 1987a,b, 1992; McKinley et al., 1991) (Fig. 19). In addition, brain nuclei localized within the BBB, such as the PVN, SON, amygdala, or NTS, showed positive labeling for ANGII receptors (Gerstberger et al., 1987a,b). Neuronal activity recorded extracellularly in vitro from hypothalamic slice preparations of the duck brain has revealed large fractions of ANGII-responsive neurons in the SFO with activation as the prevailing ANGII effect (Matsumura and Simon, 1990a). In analogy to possible inhibitory effects on salt gland function and competitive actions at the ANGII receptive sites in the SFO, angiotensin I and angiotensin I11 proved to be uneffective, with lsar-sile-ANGII fully inhibiting the ANGII-induced stimulation of neuronal discharge (Fig. 20). Adaptation of birds reared on freshwater to hypertonic saline caused hypertrophy of the supraorbital salt glands, elevated plasma and cerebrospinal fluid concentrations of ANGII (Gray and Simon, 1985, 1987; Brummermann and Simon, 1990), and an up-regulation of ANGII receptor density in the SFO exclusively at unchanged binding affinity. This might indicate that elevated endogenous ANGII reduced salt gland output of water and electrolytes via central nervous action under conditions of reduced ECFV (Gerstberger et al., 1987a). Moreover, the threshold concentration of ANGII excitation for SFO neurons was found to decrease from M ANGII in freshwater-adapted birds to M ANGII in those adapted to saline, suggesting that the salt-adapted ducks are more responsive to ANGII than freshwater birds (Simon et al., 1989; Matsumura and Simon, 1990b).

FIG. 20 Hormonal control of avian salt gland function (angiotensin 11). (A) Dose-dependent (A-C) inhibitory action of peripherally administered angiotensin I1 (ANGII) on salt gland function in the Kelp gull under conditions of steady-state osmolal excretion (dashed line). (B) Effect of ANGII on constant sodium excretion via the supraorbital salt glands of a duck in response to hypertonic saline loading (SALT) before (left) and after ganglionic blockade of secretion by mecamylamine (MECA) application (right). To reestablish steady-state secretion despite effective ganglionic blockade, the muscarinic agonist metacholine (MCH) was infused systemically. (C) Firing rate recorded from a subfomical organ (SFO) neuron in a duck hypothalamic tissue slice that was superfused with medium containing ANGII (upper left), angiotensin 111 (ANGIII) (upper right), the highly specific receptor antagonist 'sar-*ile-ANGII with ANGII (lower left), and again ANGII (lower right). [(A) From Gray and Erasmus, 1989a (modified), with permission; (B) from Butler e t a / . . 1989 (modified), with permission; (C) from Matsumura and Simon, 1990a (modified), with permission.]

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C. Atrial Natriuretic Factor There is now abundant evidence to show that the heart functions as an endocrine organ to process and secrete a peptide that plays a vital role in osmoregulation. This hormone, ANF, has been most extensively characterized by studies in mammals (DeBold et al., 1981; Genest and Cantin, 1985; Bie et al., 1988). Atrial natriuretic factor is also present in birds (Miyata et al., 1988; Toshimori et al., 1990), and its actions appear to be designed to counteract saltand fluid-conserving systems (Schiitz and Gerstberger, 1990; Gray et a[., 1991a,b; Schutz et af.,1992a). Early studies with mammalian forms of ANF failed to demonstrate any effect of this peptide on avian salt gland function (Wilson, 1987a; Langford and Holder, 1988). The successful isolation and identification of ANF from the chicken atria, however, enabled this peptide to become commercially available (Miyata et al., 1988). One study with this avian form of ANF has provided compelling evidence that ANF has a role in the regulation of salt glands. Schutz and Gerstberger (1990) showed that elevation of the circulating levels of ANF within the physiologic range by ic and/or iv infusion in ducks with actively secreting salt glands resulted in a marked enhancement of both the rate of secretion and the osmotic concentration of the fluid (Fig. 21). Preliminary data also indicated a vasodilatory effect of bird-specific ANF on the salt gland vasculature, and enhanced secretion even in the presence of ganglionic blockade, suggestive of a direct glandular ANF action in general. Moreover, receptor autoradiography of the salt glands revealed a uniform distribution of specific ANF binding sites, with a marked density throughout the parenchyma of the glands, but not in the arterioles supplying them (Fig. 21). Further receptor studies with enriched plasma membrane fractions of salt gland tissue showed that the binding sites were of a single class and high affinity. The direct action of ANF on salt gland secretion is short-lasting, and it has been suggested that this may be due to receptor-mediated internalization of the hormone (Schutz and Gerstberger, 1990). This would be consistent with the observations of Lange et al. (1989) using antibodies directed against mammalian ANF. The authors demonstrated the presence of ANF-like immunoreactivity in duck salt glands, although this could not be repeated by Schiitz and Gerstberger (1990). The transiency of the salt gland response to ANF may also indicate that systemic ANF acts as an emergency hormone, e.g., in cases of rapid vascular volume expansion resulting from postprandial salt and fluid absorption. The nonhomologous salt-secreting rectal gland of the elasmobranchs was also shown to be stimulated by ANF of mammalian sequence as well as peptidergic extracts of the shark heart (Solomon et al., 1985b). Blood flow through the gland and glandular chloride secretion were enhanced to a degree comparable to that resulting from ECFV expansion known to induce atrial ANF release in these animals. Experimental treatment of monolayer cultures of rectal gland ep-

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FIG. 21 Hormonal control of avian salt gland function (adrenal steroids and ANF). (a) Comparable time-dependent sodium excretion via the salt glands in sham-operated and adrenalectomized Pekin ducks during osmotic stimulation. (b) Stimulatory action of atrial natriuretic factor (ANF) infused into the carotid arteries of a saltwater-acclimated Pekin duck at threshold conditions of secretion on salt gland osmolal excretion. (c) Receptor autoradiogram of a duck salt gland tissue section incubated with radioiodinated bird-specific ANF ( '251-BH-labeled BH-chANF). Positive staining can be localized throughout the secretory tissue of single glandular lobes (GL). Bar, 800 pm. [(a) From Butler e t a / . , 1989 (modified), with permission; (b,c) from H. Schiitz and R. Gerstberger, Endocrinologv, 127, 1718-1726, 1990; 0The Endocrine Society.]

ithelial cells with ANF led to a pronounced rise in Cl--dependent, bumetanidesensitive SCC irrespective of the site of ANF application (luminal, abluminal) (Karnaky et al., 1991). Silva and co-workers (1987), however, reported that ANF per se proved to be ineffective in eliciting rectal glandular ion transport. These authors claim an indirect ANFergic action via release of VIP from neural stores within the gland with subsequent VIP-induced CI - secretion. In analogy to the postulated action of ANGII on brain structures outside the BBB, ANF might also modulate osmoregulatory effector systems such as the supraorbital salt gland via interaction with neurons in the SFO or organum vasculosum of the laminae terminalis (OVLT), two CVOs heavily endowed with ANF-specific binding sites (Schutz et al., 1992b). The existence of an avian brain-intrinsic ANF system in addition to that of the periphery can be derived from immunocytochemic and radioimmunologic data, and might be of importance with respect to the long-term control of salt gland function. Experiments

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using hypothalamic application of ANF by icv perfusion or local microdialysis techniques may provide valuable information concerning the role of central ANF not only in salt gland regulation but also in avian osmoregulation in general.

D. Prolactin There is conflicting evidence concerning the possible role of prolactin in salt gland control. Prolactin has been shown to partially restore salt gland function depressed by adenohypophysectomy (Ensor and Phillips, 1972; Ensor et al., 1972), and to induce marginal salt gland secretion (Peaker and Phillips, 1969; Peaker et al., 1970). The onset of salt gland secretion was also found to be much earlier in prolactin-treated birds than in those given ACTH, indicating that prolactin may have a direct effect on salt glands (Peaker et al., 1970). In contrast to these findings, further studies have shown that prolactin has no effect on salt gland secretion (Harvey and Phillips, 1982) and, moreover, blockage of prolactin secretion by bromocryptine did not alter the salt gland response to hypertonic saline loading. It has been suggested (Harvey and Phillips, 1982; Holmes and Phillips, 1985) that any effect prolactin may have upon salt gland function is indirect and probably related to its stimulation of food and water intake (Ensor, 1975, 1978). Since food or water deprivation reduced salt gland activity (Ensor and Phillips, 1972; Phillips and Harvey, 1980), the restoration of these parameters to normal in hypophysectomized birds or the maintenance of feeding in birds adapted to hypertonic saline may be the means by which prolactin indirectly influences avian salt gland function.

E. Arginine Vasotocin The role of arginine vasotocin (AVT), the avian antidiuretic hormone, in the control of renal function has been firmly established (Ames et al., 1971; Braun and Dantzler, 1972; Stallone and Braun, 1985, 1986; Gerstberger et al., 1985; Gray and Erasmus, 1988). Strong evidence indicates, however, that AVT has no, or only limited, effect on extrarenal salt gland secretion. First, neurohypophysectomized ducks, which lack AVT and demonstrate all the symptoms of diabetes insipidus, have salt glands that respond normally to hypertonic saline loading (Wright et al., 1967; Bradley and Holmes, 1971). Second, there is no consistent relationship between circulating AVT levels and salt gland activity, so that high plasma AVT concentrations can exist during conditions of both salt gland stimulation and inhibition (Gerstberger et al., 1984b,c). The effects of AVT on salt glands reported so far have not been consistent, with facilitation (Holmes and Adams, 1963; Lanthier and Sandor, 1967) and inhibition (Gill and Burford, 1969; Peaker, 1971) being observed. Intracarotide-

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ally administered AVT (20 ng/min/kg bw) did not alter ongoing steady-state salt gland secretion in the saltwater-acclimated Pekin duck, but did induce mild vasoconstriction in the salt glands at unchanged mean arterial pressure and heart rate (Gerstberger, 1991). In all cases, however, the doses of AVT required to elicit these effects resulted in supraphysiologic plasma concentrations of the hormone, thus questioning the physiologic significance of its actions (Simon, 1982; Arad and Skadhauge, 1984; Stallone and Braun, 1986; Gray and Erasmus, 1989a,b). Receptor binding studies using tritiated arginine vasopressin as radioligand did not result in positive labeling of secretory or vascular salt gland structures, but did reveal high densities of specific binding sites in both glomeruli and tubules of the collecting ducts in the avian kidney (Keil, 1990; Keil et al., 1990). Therefore AVT does not appear to play a major role in hormonal control of avian salt gland function.

F. Other Hormones There are a number of hormones that have been incidentally considered possible candidates for a role in salt gland regulation. In each case, however, it is difficult to ascribe a clear physiologic action to the hormones simply because there are insufficient data available. Three of these hormones are discussed below. 1. Thyroid Hormones

Ensor et al. (1969) demonstrated that the response of salt glands to an oral salt load is delayed by thyroidectomy. If the same load is administered iv, the salt gland functions normally. This suggests that thyroid hormone depletion affects intestinal transport rather than directly influencing salt glands. Additionally, salt glands may depend on thyroid hormones indirectly via their regulation of cell metabolism and growth (Harvey and Phillips, 1982).

2. Substance P

In one study in ducks, large doses of substance P stimulated the rate of fluid output at marginal salt gland activity, without changing its electrolyte concentration (Cheeseman et al., 1975). Conversely, Wilson (1987b) found that in the same species substance P reduced the electrolyte content of nasal fluid without affecting the rate of its secretion. To add even more confusion, in the same study substance P increased the secretion rate in glands inhibited by ANGII, without reversing the inhibitory effect of ANGII on salt excretion. The apparent ability of substance P to dissociate changes in solute composition from changes in flow rate of salt gland fluid needs to be reevaluated and could give important insights into mechanisms underlying salt gland fluid elaboration.

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3. Catecholamines The comparison of circulating epinephrine and norepinephrine concentrations between ducks maintained on freshwater and those acclimatized to hypertonic saltwater yielded similar plasma levels of epinephrine in both groups, with norepinephrine plasma levels lower in saltwater-adapted animals (Brummermann, 1988). Indications for a catecholamine-induced modulation of salt gland function should, however, not be derived from these data. Although bilateral adrenalectomy inactivated salt gland function, injections of cortisol were able to completely restore their activity without any catecholamine supplement (Phillips et al., 1961; Butler, 1980). In addition, partial depletion of catecholamines by reserpine had no effect on the salt gland response to a hypertonic saline load (Wilson and Van Pham, 1985), and the systemic application of the a-adrenergic antagonist phenoxybenzamine did not alter either resting blood flow or the proportionality between blood flow and secretion (Kaul et d., 1983). The recent demonstration that ic injections of norepinephrine markedly reduced salt gland blood flow at only moderate reduction in osmolal excretion (Gerstberger, 1991) is probably explained by a mimicking of enhanced sympathetic innervation rather than a hormonal role for catecholamine (see Section IV,D).

VIII. Stimuludecretion Coupling To elucidate the molecular events involved in the regulation of epithelial salt secretion in the avian salt gland, the intracellular pathways transferring the extracellular signals to the cellular machinery with the final induction of Na+ and CI - secretion against an enormous concentration gradient have been studied. The contributions of the major “second and third messenger” systems, namely cyclic adenosine 3‘3‘-monophosphate (CAMP),cyclic guanosine 3 ’ ,5’monophosphate (cGMP), the inositol phosphate cycle with inositol-l,4,5trisphosphate [Ins( 1,4,5)P,] and diacylglycerol (DAG), and the intracellular calcium concentration ([Ca2+],) have been investigated primarily in isolated duck salt gland cells.

A. Cyclic Adenosine 3’,5’-Monophosphate Although some early studies tended to exclude the adenylate cyclase product cAMP from being physiologically relevant in avian salt gland secretion (Peaker and Linzell, 1975; Stewart et af., 1979), a more recent investigation (Shuttleworth and Thompson, 1987) provided evidence to the contrary. Accordingly, the application of muscarinic agonists, exogenous cAMP or forskolin, an activator

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of adenylate cyclase, to duck salt gland tissue slices stimulated ouabain-sensitive oxygen consumption. The nature of the extracellular signal responsible for activation of the adenylate cyclase-cAMP system is as yet unclear with muscarinic innervation being excluded as the natural stimulus. As mentioned earlier, duck salt glands were shown to possess VIPergic innervation in addition to the postganglionic cholinergic one, and VIP stimulated glandular secretion via membrane-bound receptors (Gerstberger, 1988). Cyclic AMP has been identified as a common second messenger of VIP in various systems (Gespach et al., 1983). This coupled system also exists in the shark rectal gland (Stoff et a/., 1979; Solomon et a/., 1984a,b). Using primary cultures of avian salt gland principal cells, the cAMP analog 8-Br cyclic AMP as well as forskolin elicited a SCC that could be potentiated by theophylline, and inhibited by both furosemide and ouabain, suggestive of an intracellular pathway involving CAMP. That VIP also stimulates ion transport in these cultures in a furosemide- and theophylline-sensitive way strongly indicates that VIP represents the prime candidate using cAMP as appropriate second messenger system in the avian salt gland (Lowy and Emst, 1987; Lowy et al., 1987) (Figs. 12,22). The role of cAMP as a second messenger in avian salt gland function might be derived from data concerning the importance of cAMP in shark rectal gland function. Using an intact animal system, ECFV expansion in the dogfish S . acanthias resulted in elevated rectal gland tissue concentrations of cAMP (Erlij and Rubio, 1986). Experiments with either the isolated gland, isolated tubules, or tissue slice preparations then showed stimulated glandular vasodilation. Hyperpolarization of tubular membranes consistent with electrogenic chloride transport, enhanced ouabain binding, and oxygen consumption due to the application of cAMP could also be demonstrated. All of these effects were abolished in the presence of furosemide, indicative of a direct stimulatory action of cAMP on the furosemide-sensitive entry of Na+ into the cell (Shuttleworth and Thompson, 1980; Forrest et a/., 1983; Shuttleworth, 1983a,b). In accordance, forskolin was found to increase basolateral membrane K+ conductance in cultured rectal gland cells, thus maintaining the driving force for apical C1- extrusion. (Valentich and Forrest, 1991; Moran and Valentich, 1991). Direct interaction of cAMP with the apical chloride conductance has been discussed as an additional mode of action of this second messenger in regulating cellular Na+ and CI- secretion in the rectal gland (Greger et al., 1987a,b; Sullivan et al., 1991).

B. Cyclic Guanosine 3’,5’-Monophosphate Another cyclic nucleotide, namely cGMP, formed by guanylate cyclase activity of membrane-associated or cytosolic origine, is also able to activate the secretory process of salt glands. Cyclic GMP has been considered by some authors to be part of the mechanisms by which neural or hormonal control of avian salt

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FIG. 22 Intracellular signal transduction in freshly dissociated avian salt gland cells. (A) Salt secretion via the principal secretory cell appears to be efferently controlled by parasympathetic innervation (muscarinic) using inositol 1,4,5-trisphosphate (IP,) and intracellular Ca2+ ([Ca2+Ii) as second messengers (SM), VIP using cyclic AMP (CAMP) as SM, and ANF using cyclic GMP (cGMP) as SM. (B) Model of agonist (Ach)-induced increase in the release of Ca2+ from intracellular stores after phosphoinositol hydrolysis with generation of diacylglycerol (DG) and IP,, and extrdcellular Ca2+ influx via specific channels. The intracellular Ca2+ pool is refilled via extracellular Ca2+ influx and energy-dependent CaZ+ uptake from the cytosol. (C) Enhanced [Ca2+], due to cholinergic stimulation (CCh) is dependent on refilling of the intracellular Ca2+ pools via extracelMar Ca2+ influx. Replacement of extracellular Ca2+ after agonist-induced depletion of internal Ca stores in Ca2+-freemedium results in a transient rise in [Ca2+],itself (asterisk). (D) Oscillations of [Caz+],increase in frequency depending on the dose of the muscarinic agonist carbachol (CCh) employed. Graded elevations in baseline [Ca2+],also occur as agonist dose increases, reaching maximally elevated sustained levels of [Ca2+Ii.(E) Oscillations in [Ca2+Iielicited by the application of CCh to the cell culture system persist in the presence of the calcium channel blocker Ni2+ at slightly reduced frequency and diminished baseline [Ca2+Ii.C,D, and E represent results from single cell analysis. (F) Time-dependent phosphorylation of a 170-kDa protein by salt gland secretory cells stimulated by the muscarinic agonist CCh. [(B.C) From Stuenkel and Emst, 1990, with permission; (D,E) from Crawford e r a / . , 1991, with permission; (F) from Torchia et al., 1991, with permission.]

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glands is transduced (Stewart et al., 1979; Holmes and Phillips, 1985; Butler et al.. 1989). In salt gland slices, guanylate cyclase stimulators, such as hydroxylamine and sodium azide, stimulated ouabain-sensitive respiration as well as Na-K-ATPase activity, as did the application of cGMP itself. This action proved to be independent of Ca2+ and was not inhibited by atropine, suggestive of cGMP-induced cellular processes not related to the parasympathetic innervation of the gland (Stewart et al., 1979; Stewart and Sen, 1981). Cyclic GMP represents the second messenger that has so far been identified exclusively for ANF in mammals (Laragh and Atlas, 1988). The fact that activation of salt gland secretion by saline loading in ducks not only increased glandular tissue cGMP concentrations but also elevated the circulating level of ANF (Stewart et al., 1979; Gray et al., 1991b) makes cGMP a more than plausible candidate involved in the secretagogue effect of ANF (Fig. 22). Comparative studies showed that the ANF-induced activation of the shark rectal gland is also mediated, at least in part, by this nucleotide, although enhanced C1- secretion in the isolated rectal gland during 8-bromo-cGMP application could not be observed (Silva et al., 1987; Karnaky et al., 1991).

C. Phosphoinositols and lntracellular Calcium The idea that activation of muscarinic acetylcholine receptors in salt gland tissue increases the turnover of cellular phosphoinositides was already indicated by the early studies of Hokin and Hokin (1967). During cholinergic stimulation of salt gland slices in vitro, the incorporation of radioactive 32P into phosphatidic acid and phosphoinositides was markedly enhanced, whereas incorporation into phosphatidyl choline and ethanolamine, major phospholipids of the plasma membrane, was negligible. These observations could then be verified using dissociated duck salt gland cells, which were either prelabeled with 32P or tritiated inositol (Fisher et al., 1983; Snider et al., 1986). Accordingly, by analogy with a number of other tissues (Agranoff et al., 1984; Berridge, 1984), receptormediated breakdown of phosphatidyl-inositol 4,5-diphosphate (PIP,) led to the formation of Ins( 1,4,5,)P, and DAG. Ins( 1,4,5,)P, generation induced via muscarinic agonists proved to be sensitive to atropine (Shuttleworth, 1990; Hildebrandt and Shuttleworth, 1991b). To identify possible effects of differentiation in avian salt gland cells on inositol phosphates, Hildebrandt and Shuttleworth (199 1b) reported that upon muscarinic receptor activation both Ins( 1,4,5)P, and Ins( 1,3,4,5)P, increased to higher levels in unstressed isolated salt gland cells compared to fully differentiated ones. This suggests a possible significance of this second messenger system in cell proliferation or differentiation during the process of salt acclimation. Ins( 1,4,5)P, has been shown to induce subsequently the release of intracellularly stored Ca’+, particularly from the endoplasmic reticulum (Berridge,

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1981; Streb et al., 1983; Burgess et al., 1984; Putney, 1986; Benidge and Galione, 1988). As indicated in Fig. 22, in avian salt glands as well phosphoinositols appear to mediate Ca2+ release from intracellular stores and may also stimulate extracellular Ca2+ entry, the latter representing the essential signal for secretory activity (Shuttleworth and Thompson, 1989). The importance of a rise in [Ca2+Iifor the secretory process could be derived from tissue slice experiments, where cholinergic stimulation of the secretory process, as indicated by changes in ouabain-sensitive respiration, proved to be fully dependent on the presence of Ca2+ in the medium (Stewart et al., 1979). Using confluent cultures of salt gland cells, cellular ion transport could be induced with the Ca2+ ionophore A23187, which also elicited a SCC blockable by ouabain and furosemide (Lowy et al., 1985a,b). Agonist-induced changes in [Ca++], have been measured in a number of studies using dissociated secretory cells from duck salt glands. In the first (Snider et al., 1986) it was found that the increase in cytoplasmic Ca2+ was entirely related to extracellular Ca2+ entry. Use of a more refined method to measure Ca2+, however, demonstrated that the initial rise in [Ca2+Iioriginated from intracellular stores, followed by an increased extracellular Ca2+ influx to maintain the elevated [Ca2+Iiconcentration and to replenish depleted intracellular stores (Shuttleworth and Thompson, 1989; Stuenkel and Emst, 1990) (Fig. 22). The mobilization of intracellularly stored Ca2+ was extremely rapid and transient, reaching a peak within 2 sec and declining to basal values after 2 min in the absence of an extracellular source of the cation. In the presence of Ca2+-containing medium, however, the fall in [Ca2+], was offset by extracellular Ca2+ entry across the plasma membrane, with [Ca2+],being sustained at or near peak values while maximally stimulating concentrations of agonist are present (Shuttleworth and Thompson, 1989; Stuenkel and Emst, 1990) (Fig. 22). This pattern of Ca2+ mobilization can be demonstrated in a large variety of nonexcitable cells (Hallam and Rink, 1985; Hallam and Pearson, 1986; Merrit and Rink, 1987; Negulescu and Machen, 1988) and in each case appears dependent on stimulation of inositol lipid breakdown. Single-cell microfluorometry of changes in [Ca2+Ii revealed [Ca2+Ii oscillations to be inducible by extracellular application of the acetylcholine analog carbachol in a dose-dependent manner. Oscillation frequencies reached extraordinarily high values compared to those of other cell types (Berridge and Galione, 1988; Cobbold, 1989; Rink and Hallam, 1989), whereas the spike amplitude remained unchanged (Crawford et al., 1991) (Fig. 22). In other exocrine glands such as rat pancreatic or parotid acinar cells, [Ca2+Iioscillation frequency proved to be independent of agonist concentration (Gray 1988; Tsunoda et al., 1990). Blockade of extracellular Ca2+ entry or reduction in extracellular Ca2+ concentration reduced frequency, but not amplitude of [Ca2+Iioscillations. It could not fully inhibit the refilling of depleted intracellular calcium stores necessary for the propagation of [Ca2+Iioscillations, in-

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dicative of both Ins( 1,4,5)P,]sensitive and -insensitive intracellular calcium pools (Hildebrandt and Shuttleworth, 1991a; Crawford et al., 1991). Analysis of inositol phosphates in principal secretory cells revealed also that raising [Ca2+Iidid not cause an increase in cellular Ins(2,3,4,5,)P4 concentration, suggestive of physiologically insignificant activation of Ins( 1.4.5)P3 3-kinase (Shuttleworth and Hildebrandt, 1991).

D. Protein Phosphorylation In salt glands, the cellular events following [Ca2+Iielevation are essentially unknown, although it is to be assumed that the Ca2+-activated K+ channels of the basolateral membrane, which are thought to play a role in the secretory process, represent an important aspect of the Ca2+-regulated stimulus-secretion coupled mechanism (Richards et al., 1989). In addition, alterations in the phosphorylation status of membrane-intrinsic proteins appeared to play a major role in the regulation of cellular ion transport (Nishizuka, 1986). Accordingly protein kinase A or C (PKA, PKC) activity might also be stimulated in the avian salt gland by extracellular primary messengers resulting in the phosphorylation of various proteins, first denied in a study by Fisher and colleagues (1983). Thus, muscarinic stimulation of suspended single secretory cells of the duckling salt gland resulted in both time- and concentration-dependent increases in the phosphorylation of a 170-kDa protein blockable by the antagonist atropine (Torchia et a/., 1991) (Fig. 22). Pharmacologic experiments using phorbolesters known to activate PKC as well as PKC inhibitors indicated that muscarinic receptor activation leads to stimulation of PKC with subsequent phosphorylation of the 170-kDa membrane-intrinsic protein. The putative physiologic significance of this protein for the secretory process is supported by phosphorylation experiments with microsomal membranes revealing the ion-dependent phosphorylation of an equal-sized protein parallel with the labeling of the catalytic subunit of the Na-K-ATPase (Russo et al., 1987).

IX. Concluding Remarks Descriptive morphologic and experimental works surveyed in this article substantiate the current concept of avian salt gland function. These supraorbitally located glands represent one of the most effective organs in the vertebrate kingdom involved in the epithelial transport of ions (sodium and chloride) against a marked concentration gradient. In an orchestrated system together with the kidneys they help to maintain avian body fluid homeostasis, and marine and estuarine birds would not survive without them.

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Acknowledgments The authors highly appreciate the support, critical suggestions, and careful revision of the manuscript by Prof. Dr. S. A. Ernst, Ann Arbor, Michigan, and hof. Dr. W. Kiihnel, Liibeck, Germany. Dr. Kiihnel supplied the authors with numerous unpublished electron micrographs of avian salt gland structure, some of them being incorporated into this review. The authors are very much indebted to Prof. Dr. E. Simon and Dr. H. Schiitz for valuable discussions and support, and to Mrs. H. Holzinger for excellent photographic work and artwork.

References Abel, J. H. (1969). J . Histochem. Cytochem. 17, 570-584. Abel, J. H., and Ellis, R. A. (1966). Am. J. Anur. 118, 337-358. Addis, J. S., Menit, W. D., Mazurkiewicz, J. E., and Barmett, R. J. (1987). Cell Biochem. Funct. 5 , 135-141. Agranoff, B. W., Murphy, P., and Sequin, E. B. (1984). J. Biol. Chem. 258,2076-2078. Allen, J. C., Abel, J. H., and Takemoto, D. J. (1975). Gen. Comp. Endocrinol. 26, 209-216. Ames, E., Steven, K., and Skadhauge, E. (1971). Am. J. Physiol. 221, 1223-1228. Anderson, B. (1978). Physiol. Rev. 58, 582-603. Anderson, B., Jobin, M., and Olsson. K. (1967). Acta Physiol. Scand. 69, 29-36. Andrews, S. B., Mazurkiewicz, J. E., and Kirk, R. G. (1983). J. Cell Biol. 96, 1389-1399. Arad, Z., and Skadhauge, E. (1984). J . E.xp. Zool. 232, 707-714. Ash, R. W. (1969). Q. J . Exp. Physiol. 54, 68-79. Ash, R. W., Pearce, J. W., and Silver, A. (1969). Q. J . Exp. Physiol. 54, 281-295. Baker, P. F. (1972). In “Metabolic Pathways” (L. Hokin, ed.), Vol. 6. pp. 243-268. Academic Press, New York. Ballantyne, B., and Wood, W. G. (1969). Cytohios 4, 337-345. Bamitt, A. E., and Goertemiller, C. C. (1985). Copeiu 2,403-409. Barmett, R. J., Mazurkiewicz, J. E., and Addis, J. S. (1983). In “Methods in Enzymology” (S. Fleischer and B. Fleischer, eds.), Vol. 96, pp. 627-659. Academic Press, San Diego. Baudinette, R. V., Norman, F. I., and Roberts, J. (1982). Aust. J . Zool. 30, 407-415. Baumgarten, H. G., Holstein, A-F., and Owman, C. (1980). Z. Zellforsch. 106, 376-397. Bech, C., Rautenberg, W., May, B., and Johansen, K. (1982). J. Comp. Physiol. B 147, 71-77. Bellamy, D., and Phillips, J. G. (1966). J. Endocrinol. 36, 97-98. Bernard, C. (1865). “An Introduction to the Study of Experimental Medicine” (Eng. trans., 1957). Dover, New York. Benidge, M. J. (1981). Mol. Cell. Endocrinol. 24, 115-140. Benidge, M. J. (1984). Biochem. J. 220, 345-360. Benidge, M. J., and Galione, A. (1988). FASEB J . 2, 3074-3082. Benidge, M. J., and Oschman, J. L. (1972). “Transporting Epithelia.” Academic Press, New York. Bie, P. (1980). Physiol. Rev. 60, 961-1048. Bie, P., Wang, B. C., Leadley, R. J., and Goetz, K. (1988). Am. J. Physiol. 254, R1614169. Bonting, S. L., Caravaggio, L. L.. Canady, M. R., and Hawkins, N. M. (1964). Arch. Biochem. Biophys. 106.49-56. Borut, A., and Schmidt-Nielsen, K. (1963). Am. J . Physiol. 204, 573-581. Bradbury, M. W. B. (1985). Circ. Res. 57,213-222. Bradley, E. L., and Holmes, W. N. (1971). J . Endocrinol. 49, 437-457. Braun, E., and Dantzler, W. H. (1972). Am. J . Physiol. 222, 617-629.

AVIAN SALT GLANDS

207

Brummermann, M. (1988). Ph.D. thesis, University of Bochum, FRG. Brummermann, M., and Simon, E. (1990). J. Comp. Physiol. B 160, 127-136. Bryant, M. G., Polak. J. M., Modlin, I., Bloom, S. R., Albuquerque, R. J.. and Pearse, A. G . E. (1976). Lancet 1, 991-993. Bulger. R. E. (1963). Anat. Rec. 147, 95-127. Burford, H. J., and Bond, R. F. (1968). Experienria 24, 1086-1088. Burger, J., and Gochfeld, M. (1984). Comp. Biochem. Physiol. A 77, 103-110. Burger, J. W. (1962). Physiol. Zoo/. 35, 205-217. Burgess, G. M., Godfrey, P. P.. McKinney, J. S., Berridge, M. J.. Irvine, R. F., and Putney, J. W. (1984). Nature (London) 309.63-66. Butler, D. G. (1980). Gen. Comp. Endocrinol. 40,15-26. Butler, D. G. (1984). J. Exp. Zool. 232, 725-736. Butler, D. G. (1987). Gen. Comp. Endocrinol. 66, 171-181. Butler, D. G.. Siwanowics. H., and Puskas. D. (1989). In “Progress in Avian Osmoregulation” (A, Chadwick and M. Hughes, eds.), pp. 127-141. Leeds Philos. SOC.,Leeds. Butler, D. G., and Wilson, J. X. (1985). Comp. Biochem. Physiol. A 81, 353-358. Butler, D. G.. Youson, J. H., and Campolin, E. (1991). J. Morphol. 207, 201-210. Butt, M. M., Johnston, H. S., and Scothome, R. J. (1985). J. Anat. 141, 231-239. Carpenter, R. E., and Stafford, M. A. (1970). Condor 72, 316-324. Chance, B., Lee, C-P., Oshino, R., and Van Rossum, G. D. V. (1964). Am. J . Physiol. 206, 461-468. Cheeseman, J., Cheeseman. P., and Phillips, J. G. (1975). Gen. Conip. Endocrinol. 27, 527-530. Chipkin, S. R., Stoff, J. S., and Aronin, N. (1988a). Ann. N . Y. Acad. Sci. 527, 605-607. Chipkin. S. R., Stoff, J. S., and Aronin, N. (1988b). Peptides 9, 119-124. Claude, P., and Goodenough, D. A. (1973). J. Cell B i d . 58, 390-400. Cobbold, P. H. (1989). News Physiol. Sci. 4, 21 1-215. Cohn, A. (1903). Arch. Mikroskop. Anar. 61. Conway, G. L.. Hughes, M. R., and Moldenhauer. R. R. (1988). Cornp. Biochem. Physiol. A 91, 671-674. Cords, E. (1904). Anat. Hefte 26,49-100. Cottle, M. K. W., and Peace, J. W. (1970). Q. J. Exp. Physiol. 55, 207-212. Cowan, F. B. M. (1986). J. Microscopy 142, 87. Crawford, K. M., Stuenkel, E. L., and Emst, S. A. (1991). Am. J. Physiol. C 261, 177-184. Crowe, J. H., Nagy, K. A., and Francis. C. (1970). Am. Soc. Zool. 10, 556. DeBold, A. J., Borenstein. H. B.. Veress, A. T.. and Sonnenburg, H. (1981). Life Sci. 28, 89-94. DeRoos, R. (1961). Gen. Comp. Endocrinol. 1, 494-512. Deutsch, H., Hammel, H. T., Simon, E., and Simon-Oppermann. C. ( 1979). J. Comp. Physiol. B 129, 301-308. Deutsch, H., and Simon, E. (1980). Pjiugers Arch. 387, 1-7. Donaldson, E. M., and Holmes, W. N. (1965). J. Endocrinol. 32, 329-336. Donaldson, E. M., Holmes, W. N.. and Stachenko, J. (1965). Gen. Comp. Endocrinol. 5 , 542-551. Douglas, D. S., and Neely, S. M. (1969). Am. Zool. 9, 1095. Doyle, W. L. (1960). E q . Cell Res. 21, 386-393. Dubinsky, W. P., and Monti, L. B. (1986). Ant. J. Physiol. C 251. 721-726. Dulzetto, F. (1965). Atti. Soc. Pelorirana 11, 179-198. Dunson, W. A. (1968). Am. J. Physiol. 215, 1512-1517. Dunson, W. A. (1976). In “Biology of the Reptilia” (G. Gaus and W. R. Dawson, eds.), pp. 413-445. Academic Press, New York. Dunson, W. N., and Dunson. M. K. (1974). Am. J. Physiol. 227,430-438. Dunson, W. A., Packer, R. K., and Dunson, M. K. (1971). Science 173,437-441. Dunson, W. A., and Taub, A. M. (1967). Am. J . Physiol. 213, 975-982.

208

RUDIGER GERSTBERGER AND DAVID A. GRAY

Duvdevani, I. (1972). J . Morphol. 137, 353-364. Ellis, R. A,, and Goertemiller, C. C. (1974). Anat. Rec. 180, 285-298. Ellis, R. A,, and Goertemiller, C. C. (1976). Cytobiologie 13, 1-12. Ellis, R. A., Goertemiller. C. C., DeLellis, R. A., and Kablotsky, Y. H. (1963). Dev. Biol. 8, 286-308. Ellis, R. A., Goertemiller, C. C., and Stetson, D. L. (1977). Nature (London) 268,555-556. Ensor, D. M. (1975). Symp. Zool. Sor. (London) 35, 129-148. Ensor, D. M. (1978). “Comparative Endocrinology of Prolactin.” Chapman & Hall, London. Ensor, D. M., and Phillips, J. G. (1972). .I. Zool. (Londonj 168, 127-137. Ensor, D. M., Simons, I. M., and Phillips, J. G. (1972). J. Endocrinol. 57, xi. Ensor, D. M., Thomas, D. H., and Phillips, J. G. (1969). J. Endocrinol. 46, x. Erbe, K-H., Gerstberger, R., Gray, D. A., and Simon, E. (1988). J . Comp. Physiol. B 158, 9-17. Erlij, D., Lodenquai, S., and Rubio, R. (1980). Bull. Mt. Desert Is/.Biol. Lab. 21, 74-76. Erlij, D., and Rubio, R. (1986). J. Exp. Biol. 122, 99-112. Erlij, D., Rubio, R., and Berne, R. (1981). Bull. Mt. Desert Is/.Biol. Lab. 21, 74-76. Ernst, S. A. (1972a). J. Hisrochem. Cytochem. 20, 13-22. Ernst, S. A. (1972b). J. Histochem. Cytochem. 20.23-38. Ernst, S. A,, and Ellis, R. A. (1969). J . Cell B i d . 40, 305-321. Ernst, S. A., Goertemiller, C. C., and Ellis, R. A. (1967). Biochim. Biophys. Acta 135,682-692. Ernst, S . A., Hootman, S. R., Schreiber, J. H., and Riddle, C. V. (1981). J . Membrane Biol. 58, 101-1 14. Ernst, S. A., and Mills, J. W. (1977). J . Cell Biol. 75, 74-94. Ernst, S. A,, and Van Rossum, G. D. V. (1982). J. Physiol. 325, 333-352. Esmann, M. (1986). Biochim. Biophys. Act0 857.38-47. Esmann, M. (1988). Biochim. Biophys. Acta 940, 71-76. Esmann, M., and Skou, J. C. (1988). Biochim. Biophys. Acta 944, 344-350. Esmann, M., Watts, A., and Marsh, D. (1985). Biochemistry 24, 1386-1393. Eveloff, J., Karnaky, K. J., Silva, P., Epstein, F. H., and Kinter, W. B. (1979). J . Cell B i d . 83, 16-32. Finge, R., Krog, J., and Reite, 0. (1963). Acta Physiol. Scand. 58, 40-47. Finge, R., Schmidt-Nielsen, K., and Robinson, M. (1958a). Am. J. Physiol. 195, 321-326. Finge, R., Schmidt-Nielsen, K.. and Osaki, H. (1958b). Biol. Bull. 115, 162-171. Fawcett, D. W. (1962). Circulation 26, 1105-1132. Fisher, S. K., Hootman, S. R., Heacock, A. M., Ernst, S. A,, and Agranoff, B. W. (1983). FEES Lett. 155.43-46. Fletcher, G. L., and Holmes, W. N. (1968). J. Exp. B i d . 49, 375-391. Fletcher, G. L., Stainer, I. M., and Holmes, W. N. (1967). J . Exp. B i d . 47, 375-392. Forrest, J. N., Wang, F.. and Beyenbach, K. W. (1983). J . Clin. Invest. 72, 1163-1167. Foskett, J. K. (1987). In “Comparative Physiology of Experimental Adaptations, Adaptations to Salinity and Dehydration” (R. Kirsch and B. Lahlou, eds.), pp. 83-91. Karger, Basel. Fourman. J. (1969). J. Anal. 104, 233-239. Frizzell, R. A., Field, M., and Schultz, S. G. (1979). Am. J. Physiol. F 236, 1-8. Gassner, D., and Komnick, H. (1983). Eur. J. Cell B i d . 29, 226-235. Gaupp, E. (1888). Morphol. Jahrhuch 14,436-489. Genest, J., and Cantin, M. (1985). Endocr. Rev. 6, 107-127. Gerstberger, R. (1988). Cell Tissue Res. 252, 39-48. Gerstberger, R. (1991). J . Physiol. (London) 435, 175-186. Gerstberger, R. (1992). In “Proceedings, Twentieth Int. Congr. Ornith.,” pp. 21 14-2121. Gerstberger, R., Healy, D. P.. Hammel, H. T.,and Simon, E. (1987a). Brain Re$. 400, 165-170. Gerstberger, R., Healy, D. P., and Hammel, H. T. (1987b). Wiss. Z. Karl Marx Univ. 36, 192-194. Gerstberger, R., Healy, D. P., Hammel, H. T., and Simon, E. (1989). In “Progress in Avian Osmoregulation” (A. Chadwick and M. Hughes, eds.), pp. 13-22. Leeds Philos. SOC., Leeds.

AVIAN SALT GLANDS

209

Gerstberger, R.. Kaul, R.. Gray, D. A.. and Simon, E. (1985). Am. J. Physiol. F 248, 663-667. Gerstberger, R., Miiller, A. R.. and Simon-Oppermann, C. (1992). Prog. Brain Res. 91, 423-433. Gerstberger, R., Sann, H., and Simon, E. (1988). Am. J. Physiol. R 255, 575-582. Gerstberger, R., Simon-Oppermann, C., and Kaul, R. (1984a). J. Comp. Physiol. E 154, 449-456. Gerstberger, R., Simon, E., and Gray, D. A. (1984b). Am. J. Physiol. R 247, 1022-1028. Gerstberger, R., Gray, D. A., and Simon. E. (1984~).J . Physiol. (London) 349, 167-182. Gespach, C., Hoa, D. H. B., and Rosselin, G. (1983). Endocrinology 112, 2597-2606. Gill, J.. and Burford, H. J. (1968). J . Exp. Zool. 168, 451-454. Gill, J., and Burford, H. J. (1969). Tissue Cell 1. 497-501. Gilmore, J. P., Dietz, J., Gilmore, C., and Zucker, 1. H. (1977a). Comp. Biochern. Physiol. A 56. 121-126. Gilmore. J. P., Gilmore, C.. Dietz, J., and Zucker, I. H. (1977b). Comp. Biochem. Physiol. A 57, 119-121. Gogelein, H., Schlatter, E., and Greger, R. (1987). Pjiigers Arch. 409, 122-125. Gray, D. A,, and Erasmus, T. (1988). Am. J. Physiol. R 255, 936-939. Gray, D. A,, and Erasmus, T. (1989a). J . Comp. Physiol. E 158, 651-660. Gray, D. A,, and ErdSmuS, T. (1989b). J. Exp. Zool. 249, 138-143. Gray, D. A,, Hammel, H. T., and Simon, E. (1986). J. Comp. Physiol. E 156, 315-321. Gray, D. A., Kaul, R.. Brummermann, M., and Simon, E. (1987). Pjiigers Arch. 409, 422-426. Gray, D. A,, Schiitz, H., and Gerstberger, R. (1991a). Gen. Comp. Endorrinol. 81, 246-255. Gray, D. A., Schiitz, H., and Gerstberger, R. (1991b). Endocrinology 128, 1655-1660. Gray, D. A,, and Simon, E. (1985). Gen. Comp. Endocrinol. 60, 1-13. Gray, D. A. and Simon, E. (1987). Am. J. Physiol. R 253, 285-291. Gray, P. T. A. (1988).J . Physiol. (London) 406, 35-53. Greger. R., Gogelein, H., and Schlatter. E. (1987a). Pjliigers Arch. 409, 100-106. Greger, R., Schlatter, E., and Gogelein. H. (1987b). Pjliigers Arch. 409, 114-121. Greger, R., and Schlatter, E. (1984a). Pjliigers Arch. 402, 63-75. Greger, R., and Schlatter, E. (1984b). Pjiigers Arch. 402, 364-375. Greger, R., Schlatter, E.. and Gogelein, H. (1985). Pjiigers Arch. 403, 446-448. Greger, R., Schlatter, E., and Gogelein, H. (1986). NIPS 1, 134-136. Gumbiner. B. (1987). Am. J. Physiol. C 253, 749-758. Guth, P. H., and Leung, F. W. (1987). In “Physiology of the Gastrointestinal Tract” (L. R. Johnson, ed.), pp. 1031-1053. Raven Press, New York. Haase, P., and Fourman, J. (1970). J. Anat. 107, 382-383. Hajar, R., Sattler, F., Anderson, B. G.. and Gwinup. G. (1970). Horm. Merob. Res. 2, 35-37. Hakansson, C. H.. and Malcus, B. (1969). Acra Physiol. Scand. 76, 385-392. Hakansson, C. H., and Malcus, B. (1970). Acta Physiol. Srand. 78, 249-254. Hallam, T. J., and Pearson, J. D. (1986). FEES Lett. 207, 95-99. Hallam, T. J., and Rink, T. J. (1985). FEES Lett. 186, 175-178. Hammel, H. T. ( 198 1 ). Adv. Physiol. Sci. 18, 77-80. Hammel, H. T. (1989).In “Progress in Avian Osmoregulation” (A. Chadwick and M. Hughes, eds.), pp. 163-181. Leeds Philos. Soc.. Leeds. Hammel, H. T., and Maggert, J. E. (1983). Physiologist 26, A58. Hammel, H. T., Maggen, J. E.. Simon, E., Crawshaw, L., and Kaul, R. (1977). In “The Proceedings of the Third SCAR Symposium on Artic Biology: Adaptations within Antarctic Ecosystems” (G. A. Llano, ed.), pp. 489-500. Gulf, Houston. Hammel, H. T., Simon-Oppermann, C., and Simon. E. (1980).Am. J . Physiol. R 239,489-496. Hammel, H. T., Simon-Oppermann, C., and Simon, E. (1983). J . Comp. Physiol. E 149, 451-456. Hanwell. A,. Linzell, J. L., and Peaker. M. (1971a). J. Physiol. (London) 213, 373-387. Hanwell, A,. Linzell, J. L.. and Peaker, M. (1971b). J. Physiol. (London) 213, 389-398. Hanwell, A., Linzell, J. L., and Peaker, M. (1972). J. Physiol. (London) 226, 453-472. Hanwell, A., and Peaker, M. (1973). J . Physiol. (London) 234. 78-8OP.

210

RUDIGER GERSTBERGER AND DAVID A. GRAY

Hanwell, A,, and Peaker, M. (1975). J. Physiol. (London) 248, 193-205. Harvey, S., and Phillips, J. G. (1980). Gen. Comp. Endocrinol. 42, 334-344. Harvey, S., and Phillips, 5. G. (1982). Comp. Eiochem. Physiol. A 71, 537-546. Harvey, S., Phillips, J. G., and Rees, A. (1985). Gen. Comp. Endocrinol. 60, 210-214. Hayslett, J. P., Schon, D. A., Epstein, M., and Hogben, C. A. M. (1974). Am. J. Physiol. 226, 1188-1192. Heinroth, 0.. and Heinroth, M. (1928). “Die Vogel Mitteleuropas in allen Lebens- und Entwicklungsstufen photographisch aufgenommen und in ihrem Seelenleben bei der Aufzucht.” Behrmiiller, Berlin. Herbert, C. F. (1975). In “The Biology of Penguins” (9. Stonehouse, ed.), pp. 85-100. Macmillan & Co., London/Basingstone. Hildebrandt, J-P., and Shuttleworth, T. J. (1991a). FASEE J. 5, A1054. Hildebrandt, J-P., and Shuttleworth, T. J. (1991b). Am. J . Physiol. C 261, 210-217. Hokin, M. R. (1963). Eiochim. Eiophys. Acta 77, 108-120. Hokin, M. R. (1967). J . Gen. Physiol. 50, 2197-2209. Hokin, M. R., and Hokin, L. E. (1967). J . Gen. Physiol. 50, 793-81 1. Holmes, W. N. (1965). Arch. Anat. Micr. Morphol. Exp. 54, 491-513. Holmes, W. N. (1972). Fed. Proc. 31, 1587-1597. Holmes, W. N., and Adams, B. M. (1963). Endocrinology 73.5-10. Holmes, W.N., Butler, D. G., and Phillips, J. G. (1961). J . Endocrinol. 23, 53-61. Holmes, W. N., Lockwood, L. N., and Bradley, E. L. (1972). Gen. Comp. Endocrinol. 18, 59-68. Holmes, W. N., and Phillips, J. G. (1985). Eiol. Rev. 60, 213-256. Holmes, W. N., and Stewart, D. J. (1968). J . Exp. Eiol. 48.509-519. Holm-Rutili, I., and Berglindh, T. (1986). Am. J . Physiol. G 250, 575-580. Honda, K., Negoro, H., Dyball, R. E. J., Higuchi, T., and Takano, S. (1990). J. Physiol. (London) 431, 225-241. Hootman, S. R., and Ernst, S. A. (1980). Am. J. Physiol. C 238, 184-195. Hootman, S. R., and Ernst, S . A. (1981a). Am. J. Physiol. R 241, 77-86. Hootman, S. R., and Ernst, S. A. (1981b). J. CellEiol. 91, 781-789. Hootman, S. R., and Ernst, S. A. (1982). Am. J . Physiol. C 243. 254-261. Hootman, S. R., Ernst, S. A., and Philpott, C. W. (1987). In “Comparative Physiology of Environmental Adaptations,” (R. Kirsch and 9. Lahlou, eds.), Vol. I , pp. 26-36. Karger, Basel. Hori, T., Simon-Oppermann, C., Gray, D. A., and Simon. E. (1986). Pfliigers Arch. 407, 414-420. Hossler, F. E. (1982). Cell Tissue Res. 226, 531-540. Hossler, F. E., and Olson, K. R. (1990). J. Exp. Zoo/. 254, 237-247. Hossler, F. E., Sarras, M. P., and Allen, E. R. (1978). Cell Tissue Res. 188, 299-315. Hudson, D. M., and Lutz, P. L. (1986). Copeiu, 247-249. Hughes, M. R. (1984). Condor 86, 390-395. Hughes, M. R. (1987). J . Comp. Physiol. E 157,261-266. Hughes, M. R. (1989). In “Progress in Avian Osmoregulation” (A. Chadwick and M. Hughes, eds.), pp. 143-161. Leeds Philos. SOC.,Leeds. Jacobson, L. L. (1813). Bull. SociCtC Philomath. (Paris) 3, 267-269. Jobert, C. (1869). Ann. Sci. Naturelles 11, 349-368. Kanosue, K., Gerstberger, R., Simon-Oppermann, C., and Simon, E. (1992). Brain Res. 569. 268-274. Kanosue, K., Schmid, H., and Simon, E. (1990). Am. J . Physiol. R 258, 973-980. Karlsson, K-A., Samuelsson, B. E., and Steen, G. 0. (1974). Eur. J . Eiochem. 46, 243-258. Kamaky, K. J., Valentich, J. D., Curie, M. G., Oehlenschlager, W. F., and Kennedy, M. P. (1991). Am. J . Physiol. C 260, 1125-1130. Kaul, R.. Gerstberger, R.. Meyer, J-U., and Simon, E. (1983). J. Comp. Physiol. E 149, 457-462. Kaul, R., and Hammel, H. T. (1979). Am. J . Physiol. R 237, 355-359.

AVIAN SALT GLANDS

21 1

Keil, R. (1990). Diploma thesis, University of Tubingen, FRG. Keil, R., Gerstberger, R., and Simon. E. (1991). J . Comp. Physiol. B 161, 179-187. Keil, R., Schutz, H.. Gray, D. A.. Gerstberger. R., and Simon, E. (1990). J . Endocrinol. Invest. 13(Suppl. 2). 29 1. Kirsch, R., Humben, W., and Simmoneaux, V. (1985). In “Transport Processes, Iono- and Osmoregulation” (R. Gilles and M. Gilles-Baillien, eds.), pp. 265-277. Springer. Heidelberg. Kirschner, L. B. (1980). Am. J . Physiol. R 238, 219-223. Klingbeil. C. K. (1985a). Gen. Comp.Endocrinol. 58, 10-19. Klingbeil, C. K. (1985b). Gen. Comp. Endocrinol. 59, 382-390. Knight, C. H., and Peaker, M. (1979). J . Physiol. (London) 294, 145-151. Kolliker, A. (1860). Wiirz. Med. Zeitschrqt 1, 6. Komnick, H. (1963a). Protoplasma 56, 274-314. Komnick, H. (1963b). Protoplusma 56, 385-419. Komnick, H. (1963~).Protoplasma 56, 605-636. Komnick, H. (1964). Protoplasma 58, 96-127. Komnick, H. (1985). In “Biology of the Integument” (J. Bereiter-Hahn, A. G. Matolsky, and K. S. Richards, eds.), Vol. 2, pp. 500-5 16. Springer, Berlin. Komnick, H., and Kniprath. E. (1970). Cytohiologie 1. 228-247. Korf, H-W. (1984). J . EX^. ZOO/.232, 387-395. Korf, H-W., Simon-Oppermann, C.. and Simon, E. (1982). Cell Tissue Res. 226, 275-300. Korf. H-W., Viglietta-Panzica, C., and Panzica, G. C. (1983). Cell Tissue Res. 228, 149-163. Kiihnel, W. (1972). Z. Zellforseh. Mikros. Anat. 134, 435-438. Kiihnel, W., Burock, G., and Petry, G . (1969a). Histochemie 19, 235-247. Kiihnel, W., Petry. G., and Burock, G. (l969b). Z . Zellforsch. Mikros. Anat. 99, 560-569. Lange, W., Unger, J., Weindl, A.. and Lang, R. E. (1989). Anat. Embryo/. 179, 465-469. Langford, H. G., and Holder, K. (1988). Clin. Res. 36, 29A. Lanthier, A., and Sandor, T. (1967). Can. J . Physiol. Pharmacol. 45,925-936. Laragh, J. H.. and Atlas, S . A. (1988). Kidney Int. 34(Suppl. 25), 64-71. Leonhardt, H. (1980). In “Handbuch der Mikroskopischen Anatomie des Menschen” (A. Oksche and L. Vollrath, eds.), Vol. 4/10, pp. 177-666. Springer, Berlin. Levine, A. M., Higgins, J. A., and Barmett, R. J. (1972). J . Cell Sci. 11. 855-873. Lindemann, B., and Voute, C. (1977). In “Frog Neurobiology” (R. Linas and W. Precht, eds.), pp. 169-2 10. Springer, Heidelberg. Lingham, R. B., Stewart, D. J., and Sen, A. K. (1980). Biochim. Biophys. Actu 601, 229-234. Lowy, R. J.. Schreiber, J. H., Dawson, D. C., and Ernst, S . A. (1985a). Am. J . Physiol. C 249, 32-40. Lowy, R. J., Dawson, D. C., and Ernst, S . A. (1985b). Am. J . Physiol. C 249, 41-47. Lowy, R. J., Dawson, D. C., and Ernst, S . A. (1989). Am. J . Physiol. R 256, 1184-1191. Lowy, R. J.. and Ernst, S . A. (1987). Am. J . Physiol. C 252, 670-676. Lowy, R. J., Schreiber, J. H., and Ernst, S . A. (1987). Am. J . Physiol. R 253, 801-808. Lundberg, J. M. (1981). Actu Physiol. Scand. 112 (Suppl. 496). 1-57. Lundberg, J. M., Hedlund, B., and Bartfai, T. (1982). Nature (London) 295, 147-149. Macchi, 1. A., Phillips, J. G., and Brown, P. (1967). J . Endocrinol. 38, 319-329. Mahoney. S. A., and Jehl, J. R. (1985). Condor 87, 389-397. Marples, B. J. (1932). Proc. Zool. Soc. (London) 4, 820-844. Marshall, A. T. (1991). J . Biol. Chem. 266, 22749-22753. Marshall, A. T., and Cooper, P. D. (1987). J . Comp. Physiol. B 157, 821-828. Marshall, A. T., Hyatt, A. D., Phillips, J. G., and Condron, R. J. (1985). J . Comp. Physiol. B 156, 2 13-227. Marshall, A. T., King. P., Condron. R. J.. and Phillips, J. G. (1987). Cell Tissue Res. 248, 179-188. Marshall, A. T., and Saddlier, S . R. (1989). Cell Tissue Res. 257, 399-404.

212

RUDlGER GERSTBERGER AND DAVID A. GRAY

Martin, B. J., and Philpott, C. W. (1973). J. Exp. Zool. 186, 111-122. Martin, B. J., and Philpott, C. W. (1974). Cell Tissue Res. 150, 193-211. Marver, D., Lear, D., Marver, P., Silva, P., and Epstein, F, H. (1986). J. Membrane Eiol. 94, 205-21 5. Matsumura, K., and Simon, E. (1990a). J. Physiol. (London) 429,281-296. Matsumura, K., and Simon, E. (1990b). J . Physiol. (London) 429, 297-308. Mazurkiewicz, J. E., and Barmett, R. J. (1981). J . Cell Sci. 48, 75-88. Mazurkiewicz, J. E., Hossler, F. E., and Barmett, R. J. (1978). J. Histochem. Cyrochem. 26, 1042-1052. McArthur, P. D., and Gorman, M. L. (1978). J. Zoo/. (London) 184, 83-90. McFarland, L. Z. (1964). Nature (London) 204, 1202-1203. McFarland, L. Z., Martin, K. D., and Freedland, R. A. (1965). J . Cell Comp. Physiol. 65, 237-242. McKinley, M. J., Denton, D. A., and Weisinger, R. S. (1978). Brain Res. 141, 89-103. McKinley, M. J., McAllen. R. M.. Mendelsohn, F. A. 0..Allen, A. M., Chai. S. Y., and Oldfield, B. J. (1991). Front. Neuroendocrinol. 11, 91-127. Merchant, J. L., Papermaster, D. S., and Barmett, R. J. (1985). J . Cell Sci. 78, 233-246. Merritt, J. E., and Rink, T. I. (1987). J . Eiol. Chem. 262, 14912-14916. Meyer, J-U.. and Intaglietta, M. (19861. Ann. Eiomed. Eng. 14, 109-1 17. Mihalkovics, J. (1898). Anat. Hefre ll(10). Mikami, S. (1975). In “Brain-Endocrine Interactions. 11. The Ventricular System” (K. M. Knigge, D. E. Scott, and H. Kobayashi, eds.), pp. 80-93. Karger, Basel. Miller, R. A,, and Riddle, 0. (1943). Proc. Soc. Exp. Biol. Med. 52, 231-233. Miyata, A., Minamino, N., Kangawa, K., and Matsuo, H. (1988). Eiochem. Eiophys. Res. Commun. 155, 1330-1337. Moran, W. M., and Valentich, J. D. (1991). Am. J . Physiol. C 260, 824-831. More, N. K., and Patil, K. P. (1988). Puvo 26, 53-58. More, N. K., and Sonawane, V. D. (1988). Pavo 26, 3-10. Miiller, A. R. (1991). Diploma thesis, University of Marburg. FRG. Miiller, A. R., and Gerstberger, R. (1992). Cell Tissue Res. 268, 99-107. Negulescu, P. A,, and Machem, T. E. (1988). Am. J. Physiol. C 254, 130-140. Nicolson, S. W., and Lutz, P. L. (1989). J . Exp. Eiol. 144, 171-184. Nishizuka, Y. (1986). Science 233, 305-312. Nitzsch, C. L. (1820). Deursch Arch. Physiol. 6, 234-269. Noms, D. 0. (1985). “Vertebrate Endocrinology.” Lea & Febiger, Philadelphia. Oelrich, T. M. (1956). M i x . Puhl. Mus. Zool. Unit!.Mich. 94, 1-122. Panzica, C. G., Korf, H-W., Ramieri, G., and Viglietti-Panzica, C. (1986). CelL Tissue Res. 243, 3 17-322. Peach, M. J. (1977). Physiol. Rev. 57, 313-370. Peaker, M. (1971). J . Physiol. (London) 213, 399-410. Peaker, M. (1978). J. Physiol. (London) 276, 66P-67P. Peaker, M., and Linzell, J. L. (1975). “Salt Glands in Birds and Reptiles.” Cambridge Univ. Press, Cambridge. Peaker, M., Peaker, S. J., Hanwell, A,. and Linzell, J. L. (1973). Comp. Eiochem. Physiol. A 44, 41-46. Peaker, M., Peaker, S. J., Phillips, J. G., and Wright, A. (1971). J . Endocrinol. 50, 293-299. Peaker, M., and Phillips, J. G. (1969). J . Endocrinol. 43, ix. Peaker, M., Phillips, J. G., and Wright, A. (1970). J. Endocrinol. 47, 123-127. Peaker, M., and Stockley, S. J. (1973). Nature (London) 243, 297-298. Peaker. M., and Stockley, S. J. (1974). Experientia 30, 158-159. Perry, M. A,, Haedicke, G. J., Bulkley, G. B., Kvietys, P. R., and Granger, D. N. (1983). Proc. Soc. Exp. Eiol.Med. 177, 447-454. Phillips, J. G., and Bellamy, D. (1962). J . Endocrinol. 24, vi-vii.

AVIAN SALT GLANDS

213

Phillips, J. G., and Harvey, S. (1980). In “Avian Endocrinology” (A. Epple and M. H. Stetson, eds.), pp. 517-532. Academic Press, New York. Phillips, J. G., Holmes, W. N.. and Butler, D. G. (1961). Endocrinology 69, 958-969. Philpott, C. W., and Templeton, J. R. (1964). Anat. Rec. 148, 395-395. Pittard, J. B., and Hally, A. D. (1973). Acta Anat. 114, 303. Putney, J. W. (1979). Pharmacol. Rev. 30, 209-245. Putney, J. W. (1986). Cell Calcium 7. 1-12. Ramieri, G. (1988). E d . Soc. I t . Eiol. Sper. 64, 835-839. Richards, N. W., Lowy, R. J.. Emst, S. A.. and Dawson, D. C. (1989). J . Gen. Physiol. 93, I 171-1 194. Riddle, C. V., and Emst, S. A. (1979). J. Membrane B i d . 45, 21-35. Rink, T.J., and Hallam, T. J. (1989). Cell Calcium 10, 385-395. Roberts, J. R., and Hughes, M. R. (1984). Can. J . 2001. 62, 2142-2145. Ruch, F. E., and Hughes, M. R. (1975). Comp. Eiochem. Physiol. A 52, 21-28. Russo, J. J., Merchant, J. C., Eager, P. R., and Barmett, R. J. (1987). Cell Biochem. Funct. 5. 1-15. Sandor, T., and Fazekas, A. G. (1973). Acta Endocrinol. (Suppl). 117,251. Sandor, T., and Fazekas, A. G. (1974). Gen. Comp. Endocrinol. 22, 348-349. Sandor, T., Lamoureux, J., and Lanthier, A. (1963). Endocrinology 73, 629-636. Sandor, T., and Mehdi, A. Z. (1981). In “Advances in the Physiological Sciences” ( G . Pethes, P. Peczely, and P. Rudas, eds.), Vol. 33, pp. 331-340. Academiai Kiado, Hungary; Pergamon, London. Sandor, T., Mehdi, A. Z., and DiBattista, J. A. (1983). Can. J . Eiochem. Cell Eiol. 61, 731-743. Sandor. T., Mehdi, A. Z., and Fazekas, A. G. (1977). Gen. Comp. Endocrinol. 32, 348-359. Sarras, M. P., Rosenzweig, L. J., Addis, J. S., and Hossler, F. E. (1985). Am. J. Anat. 174. 45-60. Schildmacher, H. (1932). J. Ornithol. 80. 293-299. Schmidt-Nielsen, B. (1976). Am. J . Physiol. 230. 514-521. Schmidt-Nielsen, K. (1960). Circulation 21. 955-967. Schmidt-Nielsen, K., Jorgensen, C. B., and Osaki, H. (1958). Am. J . Physiol. 193, 101-107. Schmidt-Nielsen, K., and Kim, Y. T. (1964). Auk 81, 160-172. Schiitz, H., and Gerstberger, R. (1990). Endocrinology 127, 1718-1726. Schiitz, H., Gray, D. A., and Gerstberger, R. ( I 992a). Endocrinology 130, 678-684. Schiitz, H., Gray, D. A.. and Gerstberger, R. (1992b). f r o g . Brain Res. 91, 63-68. Schwartz, A., Lindenmayer, G. E., and Allen, J. C. (1975). Pharmacol. Rev. 27, 3-134. Scothome, R. J. (1958). Nature (London) 182, 732. Scothome, R. J. (1960). J . Anat. 94, 581. Shorofsky, S. R., Field, M., and Fozzard, H. A. (1984). J . Membrane Eiol.81, 1-8. Shuttleworth, T. J. (1983a). Am. J . Physiol. R 245, 894-900. Shuttleworth, T. J. (1983b). J. E.rp. Eiol. 103, 193-204. Shuttleworth, T. J. (1990). Eiochem. J. 266, 719-726. Shuttleworth, T. J., and Hildebrandt, J-P. (1991). FASEE J . 5, A1044. Shuttleworth, T. J., and Thompson, J. L. (1980). J . Comp. Physiol. 140, 209-216. Shuttleworth, T. J., and Thompson, J. L. (1986). J . Exp. Eiol. 125, 373-384. Shuttleworth, T. J., and Thompson, J. L. (1987). Am. J . Physiol. R 252, 428-432. Shuttleworth, T. J., and Thompson, J. L. (1989). Am. 1. Physiol. C 257, 1020-1029. Shuttleworth, T. J., and Thomdyke, M. C. (1984). Science 225, 319-321. Shuttleworth, T. J., and Wood, C. M. (1992). Am. J . Physiol. C 262. 221-228. Silva, P., Stoff, J., Field, M.. Fine, L.. Forrest, J. N.. and Epstein, F. H. (1977). Am. J . Physiol. F 233, 298-306. Silva, P., Stoff, J. S., Solomon, R. J., L e a , S.. Kniaz, D., Greger, R., and Epstein, F. H. (1987). Am. J . Physiol. F 252, 99-103. Simon, E. (1982). Comp. Eiochem. Physiol. A 71. 547-556.

214

RUDIGER GERSTBERGER AND DAVID A. GRAY

Simon, E., Eriksson, S., Gerstberger, R., Gray, D. A,, and Simon-Oppermann, C. (1987). In “Functional Morphology of Neuroendocrine Systems” (B. Scharrer, H-W. Korf, and H-G. Hartwig, eds.), pp. 37-49. Springer, Berlin. Simon, E., Gerstberger, R., and Gray, D. A. (1992). f r o g . Neurobiol. 39, 179-207. Simon, E., Gerstberger, R., Gray, D. A,, and Matsumura, K. (1989). Acta Scand. Physiol. 136(S~ppl.583). 119-129. Simon, E., and Gray, D. A. (1989). In “Progress in Avian Osmoregulation” (A. Chadwick and M. Hughes, eds.), pp. 23-40. Leeds Philos. SOC.,Leeds. Simon, E., and Gray, D. A. (1991). Am. J. Physiol. R 261, 231-238. Simon-Oppermann, C., and Gerstberger, R. (1989). In “Progress in Avian Osmoregulation” (A. Chadwick and M. Hughes, eds.), pp. 183-205. Leeds Philos. SOC.,Leeds. Simon-Oppermann, C., Gray, D. A,, Szczepanska-Sadowska, E., and Simon, E. (1984). Pfliigers Arch. 400, 151-159. Simon-Oppermann, C., and Giinther, 0. (1990). Am. J . Physiol. R 259, 294-304. Simon-Oppermann, C., Hammel, H. T., and Simon, E. (1979). Pflugers Arch. 378,213-221. Simon-Oppermann, C., and Simon, E. (1982). J. Comp. Physiol. B 146, 17-25. Simon-Oppermann, C., Simon, E., Deutsch, H., Mohring, J., and Schoun, J. (1980). Pfliigers Arch. 387, 99-106. Simons, K., and Fuller, S. D. (1985). Annu. Rev. Cell Eiol. 1, 243-288. Skadhauge, E. (1981 ). “Osmoregulation in Birds.” Springer, Heidelberg. Skou, J. C. (1965). Physiol. Rev. 45, 596-617. Skou, J. C., and Esmann, M. (1988). In “Methods in Enzymology” (S. Fleischer and B. Fleischer, eds.), Vol. 156, pp. 43-65. Academic Press, San Diego. Smith, P. L., Welsh, M. J., Stoff, J. S., and Frizzell, R. A. (1982). J . Membrane B i d . 70, 217-226. Smith, T. W. (1988). In “Methods in Enzymology” (S. Fleischer and B. Fleischer. eds.), Vol. 156, p. 46. Academic Press, San Diego. Snider, R. M., Roland, R. M., Lowy, R. J., Agranoff, B. W., and Emst, S. A. (1986). Biochirn. Eiophys. Acta 889, 216-224. Solomon, R. J., Taylor, M., Sheth, S., Silva, P., and Epstein, F. H. (1985a). Am. J . Physiol. R 248. 638-640. Solomon, R. J., Taylor, M., Dorsey, D., Silva, P., and Epstein, F. H. (1985b). Am. J . Physiol. R 249, 348-354. Solomon, R . J . , Taylor, M., Stoff. J. S., Silva, P., and Epstein, F. H. (1984a). Am. J. Physiol. R 246, 63-66. Solomon, R. J . , Taylor, M., Rosa, R., Silva, P., and Epstein, F. H. (1984b). Am. J . Physiol. R 246, 67-7 1. Sonawane, V. D. (1987). P a w 25.65-78. Spannhof, L., and Jiirss, K. (1967). Acta B i d . Med. Ger. 19, 137-144. Stainer, I. M., Ensor, D. M., Phillips, J. G., and Holmes, W. N. (1970). Comp. Eiocheni. Physiol. 37, 257-263. Stallone, J. N.. and Braun, E. (1985). Am. J. Physiol. F 249, 842-850. Stallone, J. N., and Braun, E. (1986). Am. J. Physiol. R 250, 644-657. Stewart, D. J., and Holmes, W. N. (1970). Am. J . Physiol. 219, 1819-1824. Stewart, D. J., Sax, J., Funk, R., and Sen, A. K. (1979). Am. J . Physiol. C 237, 200-204. Stewart, D. J., Semple, E. W., Swart, G. T., and Sen, A. K. (1976). Eiochim. Eiophys. Acta 419, 150-1 63. Stewart, D. J., and Sen, A. K. (1981). Am. J . Physiol. C 240,207-214. Stirling, C. E. (1976). J . Microscopy 106, 145-157. Stoff, J. S., Rosa, R., Hallac, R., Silva, P., and Epstein, F. H. (1979). Am. J . Physiol. F 237, 138-144.

AVIAN SALT GLANDS

215

Stoff, J. S . , Silva, P., Lechan, R., Solomon, R., and Epstein, F, H. (1988). Am. J . Physiol. R 255, 212-216. Streb, H., Irvine, R. F., Benidge, M. I., and Schulz, I. (1983). Nature (London) 306, 67-69. Stuenkel, E. L., and Emst, S. A. (1990). Am. J. Physiol. C 258, 289-298. Sullivan, S. K., Swamy, K., and Field, M. (1991). Am. J. Physiol. C 260, 664-669. Sundler, F.. Alumets, J., Hakanson, R., Fahrenkrug. J., and Schaffalitzky de Muckadell, 0. B. (1978). Hisrochemisfry 55, 173-176. Swaminathan, S. (1980). J . Physiol. (London) 307, 71-83. Takei, Y., Tsuneki, K., and Kobayashi, H. (1978). Cell Tissue Res. 1891, 389-404. Takemoto. D. J., Abel, J. H., and Allen, J. C. (1975). Gen. Comp. Endocrinol. 26,226-232. Technau, G. E. (1936). J. Ornirhol. 48,51 1-617. Thesleff, S.. and Schmidt-Nielsen, K. (1962). Am. J . Physiol. 203, 597-600. Thomas, D. H., and Phillips, J. G. (1975a). Gen. Conip. Endocrinol. 26, 427-439. Thomas, D. H., and Phillips, 1. G. (1975b). Gen. Comp. Endocrinol. 26, 440-450. Thomas, D. H., and Phillips, J. G. (1978). Puvo 16, 89-104. Thornton, S. N. (1986). Bruin Res. 377, 96-104. Torchia, J . , Qu, Y., Francis, J . , Pon, D. J., and Sen, A. K. (1991). Am. J . Physiol. C 261, 543-549. Toshimori, H., Toshimori, K., Minamino, N., Kangawa, K., Oura, C., Matsukura, S., and Matsuo, H. (1990). Cell Tissue Res. 259, 293-298. Tsunoda, Y., Stuenkel, E. L., and Williams, J. A. (1990). Am. J . Physiol. C 258, 147-155. Valentich. J. D., and Forrest, J. N. (1991). Am. J . Physiol. C 260, 813-823. Van Lennep, E. W., and Komnick, H. (1970). Cytoobiologie 1.47-67. Van Lennep, E. W., and Young, J. A. (1979). In “Membrane Transport in Biology” (G. Giebisch, D. C. Tosteson, and H. H. Ussing. eds.), Vol 4B, pp. 675-692. Springer-Verlag. Berlin. Van Rossum, G. D. V. (1966). Biochim. Biophys. Actu 126, 338-349. Vemey, E. B. (1947). Proc. R. Soc. Edinburgh 135, 25-106. Vigh, B. (1971). Studiu B i d . Hung. 10, 1-149. Vigh, B., and Vigh-Teichmann, 1. (1973). lnt. Rev. Cytol. 35, 189-251. Vigh-Teichmann, I., Vigh, B. and Aros, B. (1971). Z. Zellforsch. 112, 188-200. Webb, M. (1957). Aria Zool. 38,81-203. Wilson, J. X. (1987a). Con. J . Physiol. Pharmocol. 65, 1995-1999. Wilson, J. X. (1987b). Gen. Comp. Endocrinol. 67. 256-262. Wilson, J. X., and Butler, D. G. (1980). Comp.Biochem. Physiol. A 66, 583-591. Wilson. J. X., and Van Pham, D. (1985). Comp. Biochem. Physiol. C 81, 77-79. Wilson, J. X., Van Pham, D., and Tan-Wilson, H. 1. (1985). Endocrinology 117, 135-140. Wright, A., Phillips, J. G., and Huang, D. P. (1966). J . Endocrinol. 36, 249-256. Wright, A., Phillips, J. G., Peaker, M., and Peaker, S. J. (1967). In “Proceedings, Third Asia Ocean. Congr. Endocrinol..” Vol. 2, pp. 322-327. Zucker, I. H., Gilmore, C., Dietz. J., and Gilmore, J. P. (1977). Am. J. Physiol. R 232, 185-189.