Nitrogen metabolism and excretion in the mangrove killifish Rivulus marmoratus I. The influence of environmental salinity and external ammonia
Department of Zoology, University of Guelph, Guelph, Ontario, Canada N1G 2W1
*Author for correspondence (e-mail: patwrigh{at}uoguelph.ca)
Accepted 22 October 2001
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: urea, ammonia, excretion, amino acid, hyperosmotic stress, ureogenesis, ammonia detoxification, osmoregulation, ammonia exposure, mangrove killifish, Rivulus marmoratus.
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
There is little information in the literature on the effects of external salinity on nitrogen metabolism and excretion in euryhaline fish. Osmoregulating teleosts in sea water (SW) tend to have higher plasma osmolarity and urea levels than those in fresh water (FW) (Wood, 1993), but levels of urea in marine teleosts are 4070 times lower than those found in ureosmotic regulators (e.g. marine elasmobranchs) (Price and Creaser, 1967
; Payan et al., 1973
; Forster and Goldstein, 1976
). Free amino acids (FAAs) are important intracellular osmolytes in osmoconforming marine invertebrates (Pierce, 1982
; Gilles, 1987
) and, to a lesser extent, osmoregulating teleost fishes (e.g. Lasserre and Gilles, 1971
; Venkatachari, 1974
; Assem and Hanke, 1983
). FAAs can also serve as an energy source, being directly oxidized to produce ATP for osmoregulatory purposes, such as active ion transport. R. marmoratus tolerate a much wider range of salinities (0114
) than most euryhaline teleosts (034
). At very high salinities, one might expect an elevation of tissue urea and FAA levels and a compensatory depression of urea excretion (JUrea) and ammonia excretion (JAmm) to aid in the overall osmoregulatory strategy.
In addition to changes in water salinity, R. marmoratus may also be exposed to potentially toxic levels of ammonia in their natural environment. Ammonia may accumulate in the burrows from endogenous excretion by the crabs and R. marmoratus, especially during dry seasons. Fish living in extreme or variable environments, e.g. mudskippers Periophthalmodon schlosseri (Peng et al., 1998), P. cantonensis (Iwata, 1988
) and Lake Magadi tilapia Alcolapia grahami (Wood et al., 1989
), typically have a very high tolerance to ammonia. The conversion of ammonia to the less toxic urea is one mechanism of ammonia detoxification used by several ammonia-tolerant fishes, e.g. singhi catfish Heteropneustes fossilis (Saha and Ratha, 1994
), walking catfish Clarias batrachus (Saha and Das, 1999
), Lake Magadi tilapia (Randall et al., 1989
), gulf toadfish Opsanus beta (Walsh et al., 1990
) and abehaze Mugilogobius abei (Iwata et al., 2000
). Exposure of H. fossilis to external ammonia induces enzymes of the ornithineurea cycle (OUC) (Saha and Ratha, 1994
; Saha and Das, 1999
). In other species, such as mudskippers P. schlosseri, P. cantonensis and Boleophthalmus boddaerti, excess ammonia is converted to FAAs, particularly glutamate and glutamine, which are stored within the tissues (Iwata, 1988
; Peng et al., 1998
). Both ureogenesis via the OUC and stimulation of glutamine synthetase occur in ammonia-exposed abehaze (Iwata et al., 2000
). In addition, active NH4+ transport against a blood-to-water concentration gradient was reported in P. schlosseri (Randall et al., 1999
). In preliminary studies, R. marmoratus tolerated relatively high levels of external ammonia (approximately 446 µmol l1 NH3) similar to that of the ammonia-tolerant species listed above. Hence, R. marmoratus may detoxify tissue ammonia by conversion to urea, glutamine and/or glutamate.
The objectives of the present study were to investigate the ability of the mangrove killifish to modify nitrogen metabolism and excretion in order to tolerate wide changes in external salinity and ammonia levels. In the accompanying paper (Frick and Wright, 2002), we discovered the remarkable ability of R. marmoratus to release a significant amount of endogenous ammonia by volatilization (42 %) during air-exposure. In the present study, the following two predictions were tested: (i) that, during exposure to high external salinities (up to 60
), FAAs and urea will accumulate in the tissues and, consequently, that rates of ammonia and urea excretion will decrease (as a consequence of lower rates of amino acid catabolism) and (ii) that exposure to relatively high levels of external ammonia (010 mmol l1 NH4Cl) will result in a shift towards ureotelism, evident by the accumulation of urea in the tissues and an increase in the rate of urea excretion. In addition, excess ammonia will be converted into FAAs, particularly glutamate and glutamine, which will be stored in the tissues.
An additional aim of the present study was to compare nitrogen excretion between wild-caught fish and those reared under laboratory conditions. R. marmoratus are ideal fish to breed in captivity because of their small size (<2 g) and hardiness. Large numbers of clones can be produced from only a few individuals because they are hermaphroditic. A captive colony was established at the University of Guelph a decade ago, but are the physiological responses of these captive fish the same as those of their wild counterparts? To answer this question, the pattern of nitrogen excretion in laboratory-reared fish was compared with that of wild fish. A further purpose of the study was to assess the water chemistry of the natural habitat of R. marmoratus in the mangrove forests. Total ammonia levels, pH, temperature and salinity were measured in water samples collected from crab burrows at Twin Cays, Belize.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Experimental animals
Laboratory-reared fish
A colony of Rivulus marmoratus was bred from a single fish captured near Dangriga, Belize, in 1992. Eggs were collected from the parent fish and allowed to hatch and grow to maturity (approximately 0.080.15 g), at which time they were used for experimental purposes. Fish were held under the conditions described by Soto and Noakes (1994) (25°C, 16
, pH 8, 12 h:12 h L:D cycle) and were not fed for 48 h prior to experimentation to eliminate the effects of recent feeding history on nitrogen excretion measurements.
Wild-caught fish
Fish were collected from Twin Cays and Little Lagoon Cay, near Dangriga, Belize. All experiments were conducted within 4 days of capture. Prior to experimentation, fish were kept under their natural photoperiod (approximately 2 h:12 h L:D).
Experimental protocol
An initial experiment was performed under control conditions to determine the pattern of nitrogen excretion in laboratory-reared and wild-caught fish. JAmm and JUrea were measured in laboratory-reared fish (N=18) held in individual chambers containing 15 artificial SW (prepared from Instant Ocean and distilled water) (25°C, pH 8, 6 ml) for 60 h. Similar excretion measurements were performed on wild-caught fish (N=14); 15
SW was made by diluting SW with rain water (2326°C, pH 8). Water samples were collected every 6 h for a period of 36 h for measurement of JAmm and JUrea. It should be noted that, although the pH of the crab burrow water was approximately 7 (see Table 1), the experimental water was at pH 8 to ensure consistency with the rearing conditions of the laboratory-raised fish. At the higher water pH, the ammonia equilibrium will be shifted towards the non-ionic form, thereby reducing the blood-to-water ammonia partial pressure gradient compared with the situation in wild fish (Wright and Wood, 1985
).
|
Series I
R. marmoratus were exposed to FW ([Na+]1.05 mequiv l1, [Cl]
1.47 mequiv l1, [Ca2+]
5.24 mequiv l1, [Mg2+]
2.98 mequiv l1, [K+]
0.06 mequiv l1; total alkalinity
250 mg l1; total hardness
411 mg l1; determined by Xenon Laboratories, Burlington, Ontario, Canada), 15 (control), 30 and 45
SW, all at pH 8.0. Fish were subjected to a 15
change in salinity every 2 days. For example, fish in the 45
group were first transferred from 15
(control) to 30
and then held for 2 days before transfer to 45
. Water was changed daily, and water samples were collected for later determinations of ammonia and urea concentrations (on day 1, 3, 5, 7 and 9 at each salinity, N=6).
Fish exposed for 7 days to 0, 15 and 45 SW were killed for analysis of ammonia and urea tissue levels (N=5 for the 45
group and N=6 for the 0 and 15
groups). Whole fish for FAA determination were collected following 4 days of exposure to 0, 15, 30, 45 or 60
SW (N=6). All tissues were stored at 80°C for no longer than 3 weeks before analysis. It should be noted that, although sampling times in the different experiments were not always identical, the data represent an appropriate time span (i.e. 110 days) for comparisons. Tissue ammonia, urea and FAA levels were measured in whole-body samples because it was impossible to collect adequate amounts of individual tissues because of the small size of the fish and the limited number available. As total FAA levels were not significantly different between the 45 and 60
groups, only the 45
condition was included for measuring tissue urea and ammonia levels to conserve fish.
Series II
Two separate experiments involving exposure to external NH4Cl were performed: (i) a short-term exposure to 0, 1, 2, 5 and 10 mmol l1 NH4Cl during which water samples were collected every 6 h over a 48 h period (N=6); and (ii) a long-term exposure to 0 or 5 mmol l1 NH4Cl during which water samples were collected after 1, 3, 5 and 7 days of exposure (N=5).
For both experiments, fish were placed in individual chambers containing 6 ml of 16 SW at the appropriate concentrations of NH4Cl (pH 8.0). The concentration of NH3 present in 1, 2, 5 and 10 mmol l1 NH4Cl at pH 8 was calculated to be 44, 89, 223 and 446 µmol l1 NH3, respectively. Because of the high background levels of NH4Cl in the water samples, only water urea (not ammonia) concentrations were measured.
Fish in 0 or 5 mmol l1 NH4Cl were killed after 1, 4 and 10 days of exposure for analysis of tissue ammonia and urea concentrations (N=6). Both control and ammonia-exposed fish were rinsed with ammonia-free water prior to being killed. Whole fish were collected following 1 and 4 days of exposure to 0 or 5 mmol l1 NH4Cl (N=6), for determination of FAA content, and stored at 80°C, as described above.
Analytical techniques
Ammonia and urea analysis
Water ammonia levels were quantified using the Indophenol Blue method (Ivancic and Degobbis, 1984), with the following modification. Indophenol was measured at 570 nm instead of 640 nm to increase the linear range of the assay. Water urea levels were measured using the diacetyl-monoxime method (Rahmatullah and Boyde, 1980
). Ammonia and urea excretion rates (J) were expressed as µmol-N g1 h1. All spectrophotometric measurements were performed using a Perkin Elmer UV/VIS spectrophotometer (Lambda 2) (Perkin Elmer Corp., Norwalk, CT, USA).
Ammonia and urea tissue levels of whole fish were determined using the method of Chadwick and Wright (1999), except that samples were deproteinized in 10 vols of perchloric acid (8 %). The final supernatant was analyzed for ammonia concentration using an enzymatic Sigma diagnostic kit (171-C), and urea concentration was analyzed using the method of Rahmatullah and Boyde (1980
). Ammonia and urea tissue levels were expressed as mmol-N g1 wet mass.
Amino acid analysis
Whole-fish FAA levels were measured using high-performance liquid chromatography (HPLC) [Hewlett-Packard series II 1090 liquid chromatograph equipped with an ultraviolet-visible series II diode array detector (DAD), an automatic injector and a narrow-bore (20 cmx2.1 mm) reversed-phase column (AminoQuant 79916AA-572; Hewlett-Packard)]. Internal and calibration standards were prepared from individual crystalline L-amino acids to a final stock concentration of 2 mmol l1. Amino acid stock solutions were prepared in 0.1 mol l1 HCl, with the exception of glutamine, asparagine, tryptophan and taurine, which were prepared in 0.1 mmol l1 sodium acetate buffer (pH 7.2). The internal standards for primary and secondary amino acids were norvaline and azetidine 2-carboxylic acid, respectively. Primary and secondary amino acids were derivatized with o-phthaldialdehyde (OPA) and 9-fluorenylmethyl chloroformate (FMOC), respectively. Preparation and storage of OPA and FMOC reagents were as described by Barton et al. (1995).
Frozen whole fish were ground to a fine powder under liquid nitrogen and deproteinized in 500 µl of 0.5 % trifluoroacetic acid in methanol and in the presence of a known amount of the internal standards. After centrifugation (16 215 g, 4°C) for 5 min, 1 mol l1 sodium acetate and 100 mmol l1 NaOH were added, followed by centrifugation (16 215 g, 4°C) for 25 min. Concentrations of amino acids were expressed as nmol g1 wet mass.
Statistical analyses
All data are presented as means ± standard error of the mean (S.E.M.). Single-factor analyses of variance (ANOVAs) were used to examine the differences between control and treated (salinity- or ammonia-exposed) values for tissue analysis (urea and ammonia levels) and excretion rates. Amino acid data were analyzed using a General Linear models procedure using the SAS system (version 6.12; SAS Institute Inc., Cary, NC, USA). The Tukey test was used to determine whether differences were significant between treatment and control fish (P0.05). Assumptions for normality were verified by generating appropriate residual plots. Data transformations (logarithmic, square root and inverse square root) were used when appropriate to meet the above assumptions.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Wild versus laboratory-reared fish
Laboratory-reared R. marmoratus are ammoniotelic, with 7088 % of the total nitrogen excreted (JAmm+JUrea) as ammonia. Ammonia excretion remained constant over time, with no significant changes in excretion rates between time periods after the initial 12 h (Fig. 1A). Urea excretion displayed a diurnal pattern, with significantly less (75 %) urea excreted at night (00:00 to 06:00 h) than during the day (12:00 to 18:00 h) (Fig. 1B).
|
|
|
|
|
|
|
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The pattern of nitrogen excretion in wild-caught R. marmoratus was similar to that of laboratory-reared fish. Wild and laboratory-reared fish display a diurnal pattern in urea excretion, but not ammonia excretion. In addition, ammonia and urea excretion rates were altered in the same manner in the wild and laboratory-reared fish during air-exposure (Frick and Wright, 2002). Thus, we propose that the use of laboratory-raised clones in these experiments provides an accurate depiction of nitrogen excretion in this species. A diurnal variation in JUrea has also been reported in H. fossilis (Saha et al., 1988
); however, in H. fossilis, JUrea was significantly higher at night than during the day, a situation opposite to that in R. marmoratus. Other studies have reported daily variations in JAmm (Tatrai, 1981
), but these changes were attributed to variations in food intake. R. marmoratus were not fed 2 days prior to or during experimentation, and food intake is therefore not responsible for the observed pattern.
Walsh et al. (1994) reported pulsatile urea excretion in the gulf toadfish (Opsanus beta). Within a 24 h period, O. beta excrete a single burst of urea (<3 h duration) (Wood et al., 1995
), which is correlated with low levels of endogenous cortisol in the plasma (Hopkins et al., 1995
) or administration of exogenous arginine vasotocin (Perry et al., 1998
). Although R. marmoratus (not episodically) excrete urea continuously, cortisol may be implicated in daily cycles. Vijayan et al. (1996
) suggested that cortisol might contribute to the regulation of urea production in the sea raven (Hemitripterus americanus) because increased plasma cortisol levels resulted in higher levels of plasma urea (but not ammonia). As cortisol levels vary in a diurnal pattern in some teleosts (Spieler, 1979
), cortisol may be involved in the diurnal control of urea metabolism and excretion in R. marmoratus.
Changes in environmental salinity
Water salinity had a pronounced effect on the nitrogen metabolism and excretion of R. marmoratus. We predicted that acclimation to high salinities would result in the accumulation of both urea and FAAs in the tissues and a marked reduction in JUrea, and such changes were observed. Are these changes simply a reflection of an overall metabolic shut-down and/or of dehydration of the tissues under hyperosmotic stress? We think not because JAmm and tissue ammonia levels were not depressed at higher salinities and 12 out of 21 individual FAA levels were unchanged by the hypersaline environment. Thus, we propose that the observed changes in urea excretion and tissue urea and FAA concentrations are a coordinated part of a larger osmoregulatory response required in such a high-salt environment.
The twofold elevation of tissue urea levels in osmotically challenged fish (45 ) was accompanied by a significant reduction in JUrea (Fig. 3B). Isaia (1982
) found that the permeability of the gill to small non-electrolytes (e.g. urea) was lower in SW- than in FW-acclimated rainbow trout Oncorhynchus mykiss, contrary to the increase in NH4+ permeability observed in SW-acclimated teleosts (Evans et al., 1989
; Wilson and Taylor, 1992
). Gill urea permeability is dependent, in part, on urea transport proteins in the marine dogfish and toadfish and in the lake Magadi tilapia (Walsh et al., 2000
, 2001
; Fines et al., 2001
). The lipid composition of cell membranes also influences urea permeability (Lande et al., 1995
; Fines et al., 2001
) and is affected by changes in environmental salinity (Daikoku et al., 1982
). High salinities may therefore facilitate urea retention (Fig. 5) by decreasing the permeability of the gill to urea (Fig. 4), but this hypothesis requires careful testing.
We expected that JAmm would decline with increasing external salinity as a consequence of reduced amino acid catabolism and amino acid retention, assuming no change in gill permeability to ammonia. The higher JAmm at 30 compared with 0
(Fig. 3A) may relate to greater branchial NH4+ diffusion via paracellular channels typical of marine fishes (Evans et al., 1989
; Wilson and Taylor, 1992
). In addition, euryhaline fish in SW typically have a more positive gill transepithelial potential (inside relative to outside) (Potts, 1984
), which would promote the outward diffusion of NH4+ (Wright et al., 1995
). Furthermore, if NH4+ excretion is wholly or partially Na+-dependent, then changes in the availability of Na+ in the external water will affect JAmm via Na+/H+(NH4+) exchange (Claiborne and Perry, 1991
; Claiborne et al., 1999
). Yet these potential explanations still do not account for why JAmm was not highest in fish held in 45
SW. At salinities above approximately 30
, a decrease in amino acid catabolism and nitrogen excretion may be connected to amino acid retention (Table 2).
In R. marmoratus, approximately 90 % of the rise in tissue FAA levels upon acclimation to 60 SW was due to increases in levels of non-essential amino acids (Table 2), a situation similar that reported in other teleosts acclimated to SW (Huggins and Colley, 1971
; Lasserre and Gilles, 1971
; Colley et al., 1974
; Ahokas and Sorg, 1976
). The two amino acids that dominated this change were proline and taurine. The sevenfold increase in tissue proline levels has not previously been documented in a teleost. In other teleost fishes in which proline concentration was elevated during SW acclimation, the increases were less than twofold (e.g. Huggins and Colley, 1971
; Ahokas and Sorg, 1976
). Taurine, a stabilizer of cell membranes, is not found in proteins (Huxtable, 1992
) and, thus, protein catabolism would not result in increased tissue taurine levels. This may explain why tissue taurine levels do not usually increase during hyperosmotic stress (Huggins and Colley, 1971
; Ahokas and Sorg, 1976
; Deaton et al., 1984
), but decrease during hypo-osmotic stress (Lasserre and Gilles, 1971
; Vislie and Fugelli, 1975
; Fugelli and Zachariassen, 1976
). The elevation of whole-body taurine levels in R. marmoratus can only be explained by an increase in taurine synthesis (i.e. from sulphur amino acids) (King et al., 1980
).
Essential FAAs do not typically function as osmolytes because they are not endogenously synthesized and are conserved for various metabolic functions (e.g. protein synthesis) (Ballantyne and Chamberlin, 1994). However, in the present study, increases in levels of several essential FAAs occurred during acclimation to 45 and 60
SW (e.g. threonine, valine, methionine, phenylalanine, leucine and lysine). Of these FAAs, the almost threefold increase in threonine levels in fish held at 60
contributed approximately 5 % of the increase in total tissue FAA levels. Presumably, the observed changes in essential FAAs at high external salinity were as a result of protein catabolism. Overall, the increase in levels of both essential and non-essential FAAs in tissues of R. marmoratus exposed to high salinities (45 and 60
) may be important in balancing osmolyte concentrations between intra- and extracellular compartments. Moreover, the observation that total FAA levels are not significantly different between 0, 15 and 30
suggests that, over a broad range of lower salinities, the fish are not severely osmotically challenged.
Ammonia-exposure
Urea production does not play an important role in ammonia detoxification during hyperammonia stress in R. marmoratus, contrary to our predictions (Figs 6, 7). In addition, tissue levels of glutamine, glutamate and other FAAs were unchanged after 4 days of ammonia-exposure. The elevation of tissue ammonia levels on day 1 confirms that ammonia was entering the fish and implies that JAmm was initially reversed or reduced, similar to observations in other fish species (e.g. Claiborne and Evans, 1988; Iwata, 1988
; Wright, 1993
). Quite surprisingly, tissue ammonia levels were not significantly higher after 4 and 10 days of exposure to 5 mmol l1 NH4Cl (223 µmol l1 NH3) compared with controls (Fig. 7). Randall et al. (1999
) also reported that tissue ammonia levels did not increase in P. schlosseri after 6 days of exposure to 8 mmol l1 NH4Cl; however, at pH 7.2, the NH3 concentration was only 36 µmol l1. Thus, R. marmoratus are very efficient at maintaining low levels of internal ammonia without invoking ureogenesis or temporary storage of nitrogen in FAAs.
Exposure to relatively high levels of external ammonia would reverse the blood-to-water PNH3 gradient and impair excretion. Randall et al. (1999) have documented active ammonia transport in ammonia-exposed P. schlosseri, proposing that both Na+/K+(NH4+)-ATPase and Na+/H+(NH4+) exchangers are involved in branchial ammonia excretion. It is interesting to note that, when FW-acclimated killifish were exposed to 5 mmol l1 NH4Cl (well below the tolerance limit for this species), 100 % mortality (N=6) was observed after less than 24 h (N. T. Frick and P. A. Wright, unpublished observations). In contrast, all fish survived when the same or higher concentrations of NH4Cl were added to water of 15
SW. Hence, we hypothesize that Na+ is critical for survival in hyperammonia stress, possibly related to active Na+-dependent ammonia excretion.
Examination of the literature on the nitrogen metabolism and excretion of ammonia-tolerant fishes reveals two emerging patterns. In one group, a tendency towards ureotelism has been documented (see Introduction). However, the results of the present study indicate that R. marmoratus fit into the second group that do not undergo a transition to ureogenesis. The synthesis of urea is a metabolically costly process, with at least 2 mol of ATP required to synthesize 1 mol of urea (equivalent to 2 mol of NH4+) (Wood, 1993). It may be energetically more favourable to excrete ammonia actively because 1 mol of ATP would potentially eliminate 2 mol of NH4+ on the basis of the ATP hydrolysis reaction for Na+/K+-ATPase. By inhabiting such a variable and sometimes extreme environment, however, R. marmoratus are able to secure their own niche, presumably minimizing predation and competition.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Abel, D. C., Koenig, C. C. and Davis, W. P. (1987). Emersion in the mangrove forest fish Rivulus marmoratus: a unique response to hydrogen sulfide. Env. Biol. Fish. 18, 6772.
Ahokas, R. A. and Sorg, G. (1976). The effect of salinity and temperature on intracellular osmoregulation and muscle free amino acids in Fundulus diaphanus. Comp. Biochem. Physiol. 56A, 2730.
Assem, H. and Hanke, W. (1983). The significance of the amino acids during osmotic adjustment in teleost fish. I. Changes in the euryhaline Sarotherodon mossambicus. Comp. Biochem. Physiol. 74A, 531536.
Ballantyne, J. S. and Chamberlin, M. E. (1994). Regulation of cellular amino acid levels. In Cellular and Molecular Physiology of Cell Volume Regulation (ed. K. Strange), pp. 111122. Boca Raton, FL: CRC Press.
Barton, K. N., Gerrits, M. F. and Ballantyne, J. S. (1995). Effects of exercise on plasma nonesterified fatty acids and free amino acids in Arctic char (Salvelinus alpinus). J. Exp. Zool. 271, 183189.
Chadwick, T. D. and Wright, P. A. (1999). Nitrogen excretion and expression of urea cycle enzymes in the Atlantic cod (Gadus morhua L.): a comparison of early life stages with adults. J. Exp. Biol. 202, 26532662.
Claiborne, J. B., Blackston, C. R., Choe, K. P., Dawson, D. C., Harris, S. P., Mackenzie, L. A. and Morrison-Shetlar, A. I. (1999). A mechanism for branchial acid excretion in marine fish: identification of multiple Na+/H+ antiporter (NHE) isoforms in gills of two seawater teleosts. J. Exp. Biol. 202, 315324.
Claiborne, J. B. and Evans, D. H. (1988). Ammonia and acidbase balance during high ammonia exposure in a marine teleost (Myoxocephalus octodecimspinosus). J. Exp. Biol. 140, 89105.
Claiborne, J. B. and Perry, E. (1991). Acidbase transfers in the long-horn sculpin (Myoxocephalus octodecimspinosus) following exposure to 20 % seawater and low external chloride. Bull. Mt Desert Island Biol. Lab. 30, 107108.
Colley, L., Fox, F. R. and Huggins, A. K. (1974). The effects of changes in external salinity on the non-protein nitrogenous constituents of parietal muscle from Agonus cataphractus. Comp. Biochem. Physiol. 48A, 757763.
Daikoku, T., Yano, I. and Masui, M. (1982). Lipid and fatty acid compositions and their changes in the different organs and tissues of guppy, Poecilia reticulata on seawater adaptation. Comp. Biochem. Physiol. 73A, 167174.
Davis, W. P., Taylor, D. S. and Turner, B. J. (1990). Field observations of the ecology and habits of mangrove rivulus (Rivulus marmoratus), in Belize and Florida (Teleostei: Cyprinodontiformes: Rivulidae). Ichthyol. Explor. Freshwaters 1, 123134.
Deaton, L. E., Hilbish, T. J. and Koehn, R. (1984). Protein as a source of amino nitrogen during hyperosmotic volume regulation in the mussel Mytilus edulis. Physiol. Zool. 57, 609619.
Dunson, W. A. and Dunson, D. B. (1999). Factors influencing growth and survival of the killifish, Rivulus marmoratus, held inside enclosures in mangrove swamps. Copeia 3, 661668.
Evans, D. H., More, K. J. and Robbins, S. L. (1989). Modes of ammonia transport across the gill epithelium of the marine teleost fish Opsanus beta. J. Exp. Biol. 144, 339356.
Fines, G. A., Ballantyne, J. S. and Wright, P. A. (2001). Active urea transport and an unusual basolateral membrane composition in the gills of a marine elasmobranch. Am. J. Physiol. 280, R16R24.
Forster, R. P. and Goldstein, L. (1976). Intracellular osmoregulatory role of amino acids and urea in marine elasmobranchs. Am. J. Physiol. 230, 925931.
Frick, N. T. and Wright, P. A. (2002). Nitrogen metabolism and excretion in the mangrove killifish Rivulus marmoratus. II. Significant ammonia volatilization in a teleost during air-exposure. J. Exp. Biol. 205, 91100.
Fugelli, K. and Zachariassen, K. E. (1976). The distribution of taurine, gamma-aminobutyric acid and inorganic ions between plasma and erythrocytes in flounder (Platichthys flesus) as different plasma osmolalities. Comp. Biochem. Physiol. 55A, 173177.
Gilles, R. (1987). Volume regulation in cells of euryhaline invertebrates. In Cell Volume Control: Fundamental and Comparative Aspects in Animal Cells, vol. 30 (ed. A. Kleinzeller), pp. 205247. New York: Academic Press.
Hopkins, T. E., Wood, C. M. and Walsh, P. J. (1995). Interactions of cortisol and nitrogen metabolism in the ureogenic gulf toadfish Opsanus beta. J. Exp. Biol. 198, 22292235.
Huggins, A. K. and Colley, L. (1971). The changes in the non-protein nitrogenous constituents of muscle during the adaptation of the eel Anguilla anguilla L. from fresh water to sea water. Comp. Biochem. Physiol. 38B, 537541.
Huxtable, R. J. (1992). Physiological actions of taurine. Physiol. Rev. 72, 101163.
Isaia, J. (1982). Effects of environmental salinity on branchial permeability of rainbow trout, Salmo gairdneri. J. Physiol., Lond. 326, 297307.[Abstract]
Ivancic, I. and Degobbis, D. (1984). An optimal manual procedure for ammonia analysis in natural waters by the indophenol blue method. Water Res. 18, 11431147.
Iwata, K. (1988). Nitrogen metabolism in the mudskipper Periophthalmus cantonensis: changes in free amino acids and related compounds in various tissues under conditions of ammonia loading, with special reference to its high ammonia tolerance. Comp. Biochem. Physiol. 91A, 499508.
Iwata, K., Kajimura, M. and Sakamoto, T. (2000). Functional ureogenesis in the gobiid fish Mugilogobius abei. J. Exp. Biol. 203, 37033715.
King, J. C., Abel, D. C. and DiBona, D. R. (1989). Effects of salinity on chloride cells in the euryhaline cyprinodontid fish Rivulus marmoratus. Cell Tissue Res. 257, 367377.
King, P. A., Cha, C. J. and Goldstein, L. (1980). Amino acid metabolism and cell volume regulation in the little skate, Raja erinacea. I. Oxidation. J. Exp. Zool. 212, 6977.
Lande, M. B., Donovan, J. M. and Zeidel, M. L. (1995). The relationship between membrane fluidity and permeabilities to water, solutes, ammonia and protons. J. Gen. Physiol. 106, 6784.[Abstract]
Lasserre, P. and Gilles, R. (1971). Modification in the amino acid pool in the parietal muscle of two euryhaline teleosts during osmotic adjustment. Experientia 27, 14341435.[Medline]
Payan, P., Goldstein, L. and Forster, R. P. (1973). Gills and kidneys in ureosmotic regulation in euryhaline skates. Am. J. Physiol. 224, 367372.
Peng, K. W., Chew, S. F., Lim, C. B., Kuah, S. S. L., Kok, W. K. and Ip, Y. K. (1998). The mudskippers Periophthalmodon schlosseri and Boleophthalmus boddaerti can tolerate environmental NH3 concentrations of 446 and 36 µM, respectively. Fish Physiol. Biochem. 19, 5969.
Perry, S. F., Gilmour, K. M., Wood, C. M., Part, P., Laurent, P. and Walsh, P. J. (1998). The effects of arginine vasotocin and catecholamines on nitrogen excretion and the cardiorespiratory physiology of the gulf toadfish, Opsanus beta. J. Comp. Physiol. 168B, 461472.
Pierce, S. K. (1982). Invertebrate cell volume control mechanisms: a coordinated use of intracellular amino acids and inorganic ions as osmotic solute. Biol. Bull. 163, 405416.
Potts, W. T. W. (1984). Transepithelial potentials in fish gills. In Fish Physiology, vol. 10B (ed. W. S. Hoar and D. J. Randall), pp. 326388. New York: Academic Press.
Price, K. S. and Creaser, E. P. (1967). Fluctuations in two osmoregulatory components, urea and sodium chloride, of the clearnose skate, Raja eglanteria Bosc 1802. I. Upon laboratory modification of external salinites. Comp. Biochem. Physiol. 23, 6576.[Medline]
Rahmatullah, M. and Boyde, T. R. C. (1980). Improvements in the determination of urea using diacetyl monoxime; methods with and without deproteinisation. Clin. Chim. Acta 107, 39.[Medline]
Randall, D. J., Wilson, J. M., Peng, K. W., Kok, T. W. K., Kuah, S. S. L., Chew, S. F., Lam, T. J. and Ip, Y. K. (1999). The mudskipper, Periophthalmodon schlosseri, actively transports NH4+ against a concentration gradient. Am. J. Physiol. 277, R1562R1567.
Randall, D. J., Wood, C. M., Perry, S. F., Bergman, H., Malory, G. M. O., Mommsen, T. P. and Wright, P. A. (1989). Urea excretion as a strategy for survival in a fish living in a very alkaline environment. Nature 337, 165166.[Medline]
Saha, N., Chakravorty, J. and Ratha, B. K. (1988). Diurnal variation in renal and extra-renal excretion of ammonia-N and urea-N in a freshwater air-breathing teleost, Heteropneustes fossilis (Bloch). Proc. Indian Acad. Sci. 97, 529537.
Saha, N. and Das, L. (1999). Stimulation of ureogenesis in the perfused liver of an Indian air-breathing catfish, Clarias batrachus, infused with different concentrations of ammonium chloride. Fish Physiol. Biochem. 21, 303311.
Saha, N. and Ratha, B. K. (1994). Induction of ornithineurea cycle in a freshwater teleost, Heteropneustes fossilis, exposed to high concentrations of ammonium chloride. Comp. Biochem. Physiol. 108B, 315325.
Soto, C. G. and Noakes, D. L. G. (1994). Coloration and gender in the hermaphroditic fish Rivulus marmoratus Poey. Ichthyol. Explor. Freshwater 5, 7990.
Spieler, R. E. (1979). Diel rhythums of circulating prolactin, cortisol, thyoxine and triiodothyronine levels in fishes: a review. Rev. Can. Biol. 38, 301315.
Tatrai, I. (1981). Diurnal pattern of the ammonia and urea excretion of feeding and starved bream, Abramis brama L. Comp. Biochem. Physiol. 70A, 211215.
Venkatachari, S. A. T. (1974). Effect of salinity adaptation on nitrogen metabolism in the freshwater fish Tilapia mossambica. I. Tissue protein and amino acid levels. Mar. Biol. 24, 5763.
Vijayan, M. M., Mommsen, T. P., Glemet, H. C. and Moon, T. W. (1996). Metabolic effects of cortisol treatment in a marine teleost, the sea raven. J. Exp. Biol. 199, 15091514.
Vislie, T. and Fugelli, K. (1975). Cell volume regulation in flounder (Platichthys flesus) heart muscle accompanying an alteration in plasma osmolality. Comp. Biochem. Physiol. 52A, 415418.
Walsh, P. J., Danulat, E. and Mommsen, T. P. (1990). Variation in urea excretion in the gulf toadfish Opsanus beta. Mar. Biol. 106, 323328.
Walsh, P. J., Grosell, M., Goss, G. G., Bergman, H. L., Bergman, A. N., Wilson, P., Laurent, P., Alper, S. L., Smith, C. P., Kamunde, C. and Wood, C. M. (2001). Physiological and molecular characterization of urea transport by the gills of the lake Magadi tilapia (Alcolapia grahami). J. Exp. Biol. 204, 509520.
Walsh, P. J., Heitz, M. J., Campbell, C. E., Cooper, G. J., Medina, M., Wang, Y. S., Goss, G. G., Vincek, V., Wood, C. M. and Smith, C. P. (2000). Molecular characterization of a urea transporter in the gill of the gulf toadfish (Opsanus beta). J. Exp. Biol. 203, 23572364.
Walsh, P. J., Tucker, B. C. and Hopkins, T. E. (1994). Effects of confinement/crowding on ureogenesis in the gulf toadfish Opsanus beta. J. Exp. Biol. 191, 195206.
Wilson, R. W. and Taylor, E. W. (1992). Transbranchial ammonia gradients and acidbase responses to high external ammonia concentration in rainbow trout (Oncorhynchus mykiss) acclimated to different salinities. J. Exp. Biol. 166, 95112.[Abstract]
Wood, C. M. (1993). Ammonia and urea metabolism and excretion. In The Physiology of Fishes (ed. D. H. Evans), pp. 379425. Boca Raton, FL: CRC Press.
Wood, C. M., Hopkins, T. E., Hogstrand, C. and Walsh, P. J. (1995). Pulsatile urea excretion in the ureogenic toadfish Opsanus beta: an analysis of rates and routes. J. Exp. Biol. 198, 17291741.
Wood, C. M., Perry, S. F., Wright, P. A., Bergmann, H. L. and Randall, D. J. (1989). Ammonia and urea dynamics in the Lake Magadi tilapia, a ureotelic teleost fish adapted to an extremely alkaline environment. Respir. Physiol. 77, 120.[Medline]
Wright, P. A. (1993). Nitrogen excretion and enzyme pathways for ureagenesis in freshwater tilapia (Oreochromis niloticus). Physiol. Zool. 66, 881901.
Wright, P. A., Part, P. and Wood, C. M. (1995). Ammonia and urea excretion in the tidepool sculpin (Oligocottus maculosus): sites of excretion, effects of reduced salinity and mechanisms of urea transport. Fish Physiol. Biochem. 14, 111123.
Wright, P. A. and Wood, C. M. (1995). An analysis of branchial ammonia excretion in the freshwater rainbow trout: effects of environmental pH change and sodium uptake blockade. J. Exp. Biol. 114, 329353.