Ammonia and urea excretion in Lahontan cutthroat trout (Oncorhynchus clarki henshawi) adapted to the highly alkaline Pyramid Lake (pH 9.4)

Ammonia excretion in aquatic animals is strongly dependent on environmental pH (Randall and Wright, 1989). In freshwater rainbow trout (Oncorhynchus mykiss), acute exposure to alkaline water (pH9.5) results in a severe inhibition of branchial excretion caused by a reduction or reversal of the blood-to-water gradient and a blockade of Na⁺/NH₄⁺ exchange (Wright and Wood, 1985). Urea excretion increases transiently and, by 48–72h, ammonia excretion recovers by mechanisms which are as yet unknown; nevertheless, plasma ammonia levels remain greatly elevated (Wilkie and Wood, 1991). Exposure to only slightly higher pH results in death (Jordan and Lloyd, 1964; Erichsen-Jones, 1964; Murray and Ziebell, 1984; Randall and Wright, 1989), while increased water hardness at high pH increases survival of trout (Yesaki and Iwama, 1992). The remarkable tilapia of Lake Magadi, Kenya (Oreochromis alcalicus grahami), survive in extremely alkaline water (pH10.0), but do not excrete ammonia. Instead, they detoxify ammonia and synthesize urea in the liver via the ornithine–urea cycle (Randall et al. 1989; Wood et al. 1989). Clearly, ammonia excretion in water above pH9 is a challenge and fish may utilize special mechanisms to maintain nitrogen excretion in severely alkaline environments. The Lahontan cutthroat trout (Oncorhynchus clarki henshawi; Galat et al. 1985) native to Pyramid Lake, Nevada (pH9.4), not only survive at this alkaline pH, but grow to trophy size (record: 41lbs, 1925). Introductions of other salmonid species such as the rainbow trout, coho salmon (O. kisutch) and kokanee (O. nerka) have proved unsuccessful (Galat et al. 1985; Wheeler, 1987; Coleman and Johnson, 1988; Cerveri, 1990), suggesting that the Lahontan cutthroat has evolved physiological adaptations which permit it to tolerate Pyramid Lake’s extreme environment. The composition of this terminal desert lake is unusual with a salinity of 4.4‰ and total carbon dioxide, sodium, chloride, potassium and magnesium levels 5–14 times greater than local fresh water (well water; see Table 1). Historically, the Lahontan trout spawned in the fresh water of the Truckee River, the only river flowing into the lake, but the diversion of much of its flow has rendered the breeding grounds inaccessible. Natural reproduction has essentially ceased and today the fishery is maintained by artificial propagation. Spawning fish return to a man-made stream where the eggs and sperm are air-stripped and dry-fertilized. The resulting fry are raised for approximately 12 months in well water in a hatchery. At 1 year of age, these fish are acutely transferred to ponds supplied with lake water to acclimate them for several weeks, prior to being stocked into the lake.

The objective of our study was to determine how nitrogenous wastes are excreted in Lahontan cutthroat trout adapted to Pyramid Lake water at pH9.4. In the first experiment, we measured the resting rates of ammonia and urea excretion via gills and kidney to test (a) whether absolute N-production rates were typical of salmonids; (b) whether a greater than normal proportion was excreted as urea; and (c) whether a greater than normal proportion was excreted by the kidney. The collection of urine in this experiment allowed analysis of urinary ionic composition, of interest in view of the reported abnormal histology of the kidney in Pyramid Lake trout (Galat et al. 1985). In the next set of experiments, we evaluated the possible role of the Na⁺/NH₄⁺ exchange mechanism in branchial ammonia excretion by exposing trout to the drug amiloride, a competitive Na⁺ uptake blocker (Kirschner et al. 1973; Wright and Wood, 1985). In view of the Na⁺-rich environment in which amiloride was used, we employed low-Na⁺ artificial lake water to confirm the result of this experiment. We also employed the carbonic anhydrase blocker acetazolamide to test whether acidification of the gill water boundary layer by CO₂ excretion played any role in branchial ammonia efflux (Wright et al. 1989). In the final series of experiments, we directly investigated the importance of NH₃ diffusion. The blood-to-water gradients were experimentally manipulated by acutely exposing trout to lake water of higher (pH10) and lower pH (pH7.6), and by holding fish in a ‘closed system’ for 8h in which water increased progressively.

All experiments were performed at the lakeside laboratory of the Pyramid Lake Fisheries, which kindly donated the fish and facilities used in this study. Lahontan cutthroat trout (Oncorhynchus clarki henshawi; 100–281g) had been raised for approximately 1 year in well water (pH8.4; Table 1) and then transferred to large outdoor holding ponds served with flowing lake water (pH9.4; Table 1) 3–5 weeks prior to the present study, which was conducted in May, 1991. In the laboratory, the fish were held for an additional 5–10 days without food in flowing lake water at seasonal temperature (8–12°C). One day prior to experimentation, the animals were transferred to individual darkened acrylic flux chambers (McDonald and Rogano, 1986) supplied with lake water of the same temperature at 0.5 l min⁻¹, which kept water ammonia levels below 5 µ,mol l⁻¹. Mixing and air-saturated conditions were maintained by continuous bubbling of the perimeter of the flux box with air.

For some experiments, fish were surgically fitted with dorsal aortic (DA) (Soivio et al. 1972) or internal urinary bladder (Curtis and Wood, 1991) catheters while under MS-222 anesthesia. These fish were left to recover for 24–48h in their individual chambers prior to experimentation. Urine was collected into covered vials using a siphon of -300Pa. Urine was analyzed for ammonia, urea and a range of electrolytes. Blood samples (300 µ,l) were drawn anaerobically from the DA cannula and replaced with an equal volume of heparinized Cortland saline [Wolf, 1963; sodium heparin (Sigma); 100i.u.ml⁻¹]. Blood was analysed for pH (pHe), total plasma CO₂ content and total plasma ammonia (T_amm) and urea content (T_urea). At the time of blood sampling, the transepithelial potential (TEP) between body fluids and water was also determined. Three groups of experiments were performed.

Trout (N=11) were fitted with urinary bladder catheters to allow separate measurement of branchial and renal fluxes (Wood, 1988). At the start of the experiment, inflow was stopped and water volume was reduced to approximately 3l so as to maximize the sensitivity with which changes in water ammonia and urea could be measured. Blank tests demonstrated that there was no loss of ammonia by volatilization to the air. Water samples (10ml) were taken at the start and every subsequent hour over a 3h period. Urine flow was collected over a 6–12h period. Samples were immediately frozen (-20°C) for analysis within 4 days.

In these experiments, ammonia excretion was measured in fish over a 3h control period followed by a 3h experimental period. The control period was the same as described above (group 1 experiments), after which the box was flushed with fresh acclimation water and then changed to the experimental medium. This caused no apparent disturbance to the fish; changeover was complete within 5min. Water samples (10ml) were collected at the start of each 3h period and every subsequent hour for analysis of ammonia levels. Water pH was continuously monitored and maintained at pH9.4 by adding small volumes of 1moll⁻¹KOH.

Four experiments were performed. In experiment i (amiloride), the fish were fitted with DA catheters; in experiments ii–iv, the fish were uncatheterized.

• Fish (N=9) were exposed in the external water to the drug amiloride (C₆H₈ClN₇O: Sigma, 10⁻⁴ mol l⁻¹), a sodium transport blocker, to determine the effects on ammonia excretion. Arterial blood samples and TEP measurements were taken at 0.5h and 2.5h of each 3h period. Amiloride was relatively insoluble in the alkaline lake water and it was necessary to add 0.4% ethanol to achieve complete solubility.

• A control experiment was carried out in which fish (N=6) were exposed to lake water containing 0.4% ethanol.

• Fish (N=8) were exposed to nominally sodium-free artificial lake water (low-Na⁺ water) prepared by reconstituting deionized tapwater with salts to match approximately the composition of lake water. Sodium chloride was replaced with choline chloride. The measured composition of this low-Na⁺ media is shown in Table 1. In spite of extensive flushing, it was impossible completely to eliminate the presence of Na⁺ during the experimental period. Measured water Na⁺ levels were 2.18±0.74mmol l⁻¹at the start and 2.24±0.78mmol l⁻¹at the end of the 3h test.

• Fish (N=13) were exposed to the drug acetazolamide (C₄H₆N₄O₃S₂: Sigma, 1.6X10⁻³ mol l⁻¹), a specific inhibitor of carbonic anhydrase, in lake water.

Three experiments were performed to determine the effects of changing water pH and blood-to-water ammonia gradients on ammonia and urea excretion rates.

• Changes in ammonia and urea excretion rates were measured in uncatheterized fish (N=5) exposed to lake water of pH10 prepared by titration with 1mol l⁻¹ KOH, which raised the water K⁺ level to approximately 13mmol l⁻¹. A 3h control period was followed by a 3h experimental period (pH10 water) and then a second 3h control period (i.e. recovery). Water sampling and flushing of the experimental chambers were performed as described above. Blank tests again demonstrated that losses of ammonia by volatilization over the 3h period were negligible, despite the high pH. As a precaution, water samples were acidified below pH7.0 immediately after collection, then frozen and later analyzed for ammonia and urea levels. Water pH was continuously monitored (mean=9.97±0.01) and adjusted, as necessary, with 1mol l⁻¹ KOH.

• Changes in ammonia and urea excretion rates and various blood variables were measured in cannulated fish (N=6) exposed to lake water of pH7.6. The experimental protocol was the same as described above for exposure to pH10 water, except that arterial blood samples and TEP measurements were taken at 0.5h and 2.5h of each 3h period. To avoid any possibility of hypercapnia in view of the high total CO ₂ content of lake water, the experimental water was first decarbonated by acidification to pH4.0 with 6mol l⁻¹ HCl and vigorous overnight aeration, then titrated back to experimental pH with 1mol l⁻¹ KOH before introduction into the fish chambers. This reduced water total CO₂ levels to less than 1mmol l⁻¹ and raised Cl⁻ and K⁺ levels to about 83 and 7mmol l⁻¹ respectively. Water pH was continuously monitored (mean=7.63±0.06) and adjusted with 1 mol l⁻¹ HCl and 1mol l⁻¹ KOH.

• Changes in blood and water variables were monitored in cannulated fish (N=7) exposed to gradually increasing water ammonia and urea levels. This was accomplished by holding fish in a ‘closed system’ over an 8h period without flushing the box with fresh water. Water samples were collected hourly and blood samples and TEP measurements were taken at 0, 2, 4, 6 and 8h. Water pH was continuously monitored (mean=9.39±0.01) and adjusted with 1mol l⁻¹ KOH and 1mol l⁻¹ HCl.

Water pH was monitored with a glass combination electrode (Radiometer GK2401C) coupled to a Radiometer pHM 84 acid–base analyser. Water and urine ammonia levels were measured by means of the salicylate–hypochlorite assay (Verdouw et al. 1978) and urea levels by the diacetyl–monoxime method (Crocker, 1967) using commercial reagents (Sigma 535A). The uricase phosphotungstate assay (Henry et al. 1957) with commercial reagents (Sigma 680) was employed to check for the presence of uric acid in water. Urine cations were measured by atomic absorption and anions by HPLC using methods identical to those of Curtis and Wood (1991).

Whole-blood pH (pHe) was measured with a Radiometer E5021a ‘gun’ microelectrode regulated to the experimental temperature. Plasma was separated by centrifugation (2min at 13000 g). Plasma was determined with a Corning digital CO₂ analyzer. Total plasma ammonia (T_amm) was assayed enzymatically with a diagnostic kit (Sigma 170A); total plasma urea (T_urea-N) was determined as above (Sigma 535A). Plasma urea values are expressed as T_urea-N to account for two nitrogen molecules per urea molecule.

TEP between arterial blood and the environment was determined by means of 3mol l⁻¹ KCl–agar bridges connected via Ag/AgCl electrodes to a high-impedance voltmeter (Perry and Wood, 1985). The reference electrode was placed in the water in the fish chamber and the measurement electrode was connected to the DA cannula. The system was zeroed by initially placing both electrode KCl agar tips in the water. The offset voltage was less than 2mV.

The blood-to-water partial pressure gradient for NH₃ (Ll⁠) was calculated from plasma and water total ammonia concentrations and pH using appropriate constants from Cameron and Heisler (1983) and equations outlined in Wright and Wood (1985), with one exception. In the present study, plasma levels were calculated from arterial values, whereas, in our previous study (Wright and Wood, 1985), arterial values were adjusted to account for higher ammonia levels in the venous blood. The blood-to-water electrochemical gradient for NH₄⁺ (ΔNH₄⁺) was calculated from plasma and blood NH₄⁺ concentrations (see Wright and Wood, 1985) and TEP, according to the following equation:

where

and R, F, T and Z have their usual meaning.

Net fluxes of ammonia (J_amm) and urea (J_urea) were calculated as:

where T_i and T_f refer to initial and final concentrations of water ammonia or urea in µ,mol l⁻¹, V is the volume of the system in ml (corrected for sampling deficits), t is the elapsed time in h and W is the fish mass in kg. While J_amm was measured over 1 h intervals, J_urea was determined over 2 or 3h because changes in water T_urea were not large enough to be detected reliably over 1h.

Renal flux rates of ammonia, urea and electrolytes were calculated as the product of urine flow rate (UFR) and urinary concentration.

Values are reported as means ± 1 S.E.M. (N), where N represents the number of fish employed in the experiment. Student’s two-tailed paired t-tests were used to evaluate the significance of differences between mean values (P<0.05).

Table 2 compares UFR and urinary excretion rates of Lahontan cuthroat trout in Pyramid Lake water with typical values for rainbow trout in fresh water, obtained with identical methods by Curtis and Wood (1991). Relative to rainbow trout, Lahontan cutthroat trout had lower UFR values and much higher excretion rates for most electrolytes, but especially ammonia, Na⁺, Cl⁻, Mg²⁺, SO₄²⁻, NO₃⁻ and PO₄²⁻.

Assuming that the sum of ammonia-N and urea-N represents total N-excretion in Lahontan cutthroat trout, overall rates were typically 100–200 µ,molkg⁻¹ h⁻¹ (Fig. 1). The excretion of uric acid, if it occurred, was undetectable (i.e. <15 µ,mol N kg⁻¹ h⁻¹). The majority of N-waste was excreted across the gills as ammonia (56%, Fig. 1) but renal ammonia excretion was also important (10%). Urea-N excretion across the gills also accounted for a very significant portion (32%), while renal urea-N efflux contributed an additional 2%.

J_amm did not vary significantly over 3h in either control or experimental periods; therefore, average values for each 3h period are presented in Table 3.

It was necessary to pre-dissolve amiloride in ethanol, resulting in a final ethanol concentration of 0.4% in the experimental water. Therefore, initial control experiments were performed to test the effects of ethanol alone. Ammonia excretion was unchanged in control fish exposed to trace levels of ethanol (0.4%) in lake water (Table 3). Amiloride (10⁻⁴ mol l⁻¹), a Na⁺ uptake blocker, had no effect on ammonia excretion when added to the water in the experimental period. Furthermore, ammonia excretion was unaffected by exposure to low-Na⁺ lake water, indicating that Na⁺-dependent ammonia excretion (Na⁺/NH₄+ exchange) is not an important component of net ammonia excretion in Lahontan cutthroat trout (Table 3).

Blood measurements made during the amiloride experiment (temperature=9°C) are summarized in Table 4. Blood pH was about 8.15, plasma about 8mmol l⁻¹, plasma T_amm between 190 and 270 µ,mol l⁻¹, plasma T_urea-N about 30-to 50-fold greater than T_amm, and TEP slightly negative (about -5 mV) with respect to the environmental water. Plasma T_amm increased significantly during both control and experimental periods. None of these variables was affected by the amiloride treatment.

Acetazolamide (1.6X10⁻³ moll⁻¹), a carbonic anhydrase inhibitor, was added to the water in the experimental period to test the importance of acidification of the gill water boundary layer. If carbonic-anhydrase-dependent CO₂ hydration in the gill epithelial boundary layer significantly lowers water pH near the gill surface, then acetazolamide should reduce acidification and inhibit ammonia excretion, at least temporarily. However, ammonia excretion was unchanged by the acetazolamide treatment, indicating that acidification of the boundary layer is not critical to the maintenance of ammonia excretion rates (Table 3).

If ammonia excretion is dependent on passive diffusion of NH3, then one would predict that acute changes in external water pH would alter the rate of ammonia excretion. The ammonia excretion rate was significantly decreased (-52%) over 3h of exposure to pH10 water compared to the control value (Fig. 2A). When rates were compared on an hourly basis, however, the inhibition of ammonia excretion was significantly different from the control value only in the final hour. Ammonia excretion returned to control rates in the recovery period. Urea excretion was unaffected by the experimental treatment (Fig. 2B).

Ammonia excretion increased threefold in response to exposure to pH7.6 water, with the greatest stimulation occurring during the first hour (Fig. 2C). In the subsequent recovery period, ammonia excretion returned to control rates. Urea excretion was unaffected by pH7.6 water (Fig. 2D).

Plasma T_amm decreased significantly during the experimental period, while water T_amm increased as a result of enhanced ammonia excretion rates (Fig. 3). The blood-to-water gradient at 0.5 and 2.5h during the experimental period was unchanged. The estimated gradient (Fig. 4, see below) at 0h, however, may give a better indication of the immediate effects of pH7.6 water. The electrochemical gradient for NH₄+ (ΔNH₄+) was significantly decreased during pH7.6 exposure, with negative values in the last hour of the experimental period, indicating a reversal of the gradient (Fig. 3).

Fish became slightly acidotic during exposure to pH7.6 water (temperature=9.5°C), as pHe decreased by about 0.15pHunits (Table 5). Total levels decreased by approximately 1mmol l⁻¹. During the recovery period, pH and were not significantly different from control values. TEP became more negative by about -4 mV during exposure to pH7.6; it returned to control values during recovery.

Fig. 4 superimposes the calculated blood-to-water gradient on the hourly ammonia excretion rates. The gradients at 0h of the experimental and recovery period were estimated because it was apparent that the gradient at 0.5h (Figs 3 and 4) was not representative of the gradient immediately after exposure to water of a different pH. For example, blood samples taken 0.5h into the experimental period were already depleted of ammonia (Fig. 3) because of the rapid loss from the blood into the water. The estimated gradient at 0h of the experimental and recovery periods was conservatively calculated using the plasma level at 2.5h of the preceding period and the water value at 0h of the period in question. The estimated gradient at 0h of the experimental period (24.3±3.3mPa) is probably more representative of the initial gradient upon exposure to pH7.6 water, compared to the gradient at time 0.5 h (7.5±0.9mPa). Similarly, the estimated gradient at 0h of the recovery period is lower (-5.6±2.1mPa) and therefore probably more realistic than the value at 0.5 h (3.1±1.9mPa), in view of the low plasma T_amm at the end of the experimental period (Fig. 4). Changes in ammonia excretion rate appear to be associated with changes in the gradient when fish were initially exposed to pH7.6 water and recovery water (pH9.4). Between 0.5 and 2.5h of the experimental period, however, J_amm was significantly elevated despite a return of the gradient to control values. It may be that carrier-mediated transport mechanisms were also involved in stimulating ammonia excretion during exposure to pH7.6 water.

Ammonia excretion exhibited a step-wise decrease over the first 6h when fish were held in a ‘closed system’ (Fig. 5). From 6 to 8h, however, there was a sharp increase in ammonia excretion rates to levels comparable to initial excretion values (0–1h). As in other experimental treatments, urea excretion rates were not significantly changed (Fig. 5).

Plasma T_amm increased in a linear manner, from 183±17 µ,mol l⁻¹ at 0h to 716±106 µ,mol l⁻¹ after 8h (Fig. 6). Likewise, water T_amm increased linearly, but accumulation after 8h was only 56±6 µ,mol l⁻¹. Plasma and water levels increased in parallel over the experimental treatment (Fig. 6). There was a small, progressive decrease in the electrochemical gradient for NH₄+ over the 8h period from 79.2±1.6mV to 63.2±3.9mV.

There was a significant blood acidosis in the first 4h of the experiment (temperature 10°C; Table 6). Plasma was not significantly affected by the experimental treatment. TEP increased very slightly at 6h, but was not significantly different from the initial value by 8h.

Fig. 7 illustrates the blood-to-water gradient and ammonia excretion rates. Plasma levels were calculated based on blood samples collected at 0, 2, 4, 6 and 8h. Hourly plasma levels were estimated by linear interpolation. Over the first 5h of the ‘closed system’ experiment, the gradient and ammonia excretion rates both decreased over time. The marked increase in J_amm after 6h was associated with an increase in the mean gradient, but the data were highly variable.

Most teleost fish excrete the majority of their nitrogen wastes as ammonia across the gills. Acute exposure to alkaline water (pH9.5) in rainbow trout inhibits ammonia excretion (Wright and Wood, 1985). One possible way to maintain nitrogen excretion in alkaline water would be to increase the proportion of nitrogen wastes excreted as urea. In freshwater rainbow trout, urea excretion accounts for approximately 13% of total nitrogen excretion, with the remainder being excreted as ammonia (Olson and Fromm, 1971; Wilkie and Wood, 1991). In contrast, the tilapia of Lake Magadi, Oreochromis alcalicus grahami, excrete 100% of their nitrogen wastes as urea, presumably an adaptation for survival in water of pH10 (Randall et al. 1989; Wood et al. 1989). Lahontan cutthroat trout adapted to pH9.4 water excreted 34% of their nitrogen wastes as urea (Fig. 1), whereas those adapted to neutral water excreted only 10% of their nitrogen wastes as urea (M. Wilkie, G. K. Iwama, P. A. Wright and C. M. Wood, in preparation). It is noteworthy that the absolute rates of urea excretion in the Lahontan cutthroat trout at pH9.4 were very similar to those reported by Wilkie and Wood (1991) for rainbow trout in neutral water. The relative rate of urea versus ammonia excretion, however, was much higher in the cutthroat trout because the absolute rate of ammonia excretion was about 50% lower compared to rates in the rainbow trout and about 70% lower compared to rates in Lahontan cutthroat trout acclimated to well water (M. Wilkie, G. K. Iwama, P. A. Wright and C. M. Wood, in preparation). Hence, one strategy for effectively excreting N-wastes in the alkaline waters of Pyramid Lake is to decrease the rate of ammonia excretion and to increase the relative proportion of nitrogen wastes excreted as urea. Urea excretion rate, however, was not regulated with acute changes in water pH (Fig. 2) or elevated water ammonia and urea levels (Fig. 5), although it increased significantly during longer-term exposure to pH10 in the accompanying study (Wilkie et al. 1993).

The pathway for urea synthesis in the tilapia O. a. grahami (Randall et al. 1989), the toadfish Opsanus beta (Read, 1971; Mommsen and Walsh, 1989), the freshwater air-breathing teleost Heteropneustes fossilis (Saha and Ratha, 1986) and higher vertebrates is the ornithine–urea cycle. The majority of teleosts, however, synthesize urea at relatively low rates through the metabolism of purines (uricolysis) or arginine. In the accompanying study, we have shown that Lahontan cutthroat trout are typical of other teleosts in that they have low activities of the ornithine–urea cycle enzymes but significant activities of uricase, allantoicase and allantoinase, the three enzymes involved in uricolysis (Wilkie et al. 1993).

In the freshwater rainbow trout, renal ammonia excretion accounts for only about 2 % of total ammonia excretion, with the remainder excreted across the gills (McDonald and Wood, 1981; Wood, 1988; Curtis and Wood, 1991). In the present study, the renal component of ammonia excretion in Lahontan cutthroat trout was 14% (Fig. 1; 10% of total nitrogen excretion), a significantly greater value than that of the rainbow trout. Therefore, the greater role of the kidneys in ammonia excretion in Lahontan cutthroat trout on both an absolute and relative basis (see Table 2) may be another adaptation to maintain nitrogen excretion in a severely alkaline environment. Further work will be required to determine whether this difference results from a modification of active (e.g. ionic secretion, renal ammoniagenesis) or passive (e.g. filtration, diffusive secretion) mechanisms for renal ammonia excretion. Interestingly, there appears to be no accompanying modification of renal urea excretion (Table 2).

The lower UFR of Lahontan cutthroat trout relative to rainbow trout in fresh water (Table 2) clearly reflected the lower osmotic gradient for water entry at the gills in Pyramid Lake water, which is equivalent to about 13% sea water (Table 1). Despite this lower UFR, the kidney made a much larger contribution to the excretion of most ions (Table 2). These higher excretion rates (for Na⁺, Cl⁻ and Mg²⁺) reflected the unusual saline conditions of Pyramid Lake, in which levels of all three ions are markedly elevated (Table 1), whereas the more normal excretion rates for Ca²⁺ and K⁺ were correlated with the relatively low levels of these ions in the water. However, the higher rates of phosphate and NO₃⁻ excretion in the urine of Lahontan trout did not reflect water composition, for levels of these electrolytes were below detection in Pyramid Lake water. Finally, the higher SO₄²⁻ excretion is noteworthy, because Galat et al. (1985) correlated water SO₄²⁻ concentration with glomerular swelling, congestion and deposition of hyalin in the kidney tubules in Lahontan trout sampled from Pyramid and other alkaline lakes in the western United States. SO₄²⁻concentration was only moderately elevated in Pyramid Lake water relative to most standard fresh waters (Table 1).

In most teleosts, branchial ammonia excretion depends largely on the transepithelial NH₃ gradient (deVooys, 1968; Kormanik and Cameron, 1981; Cameron and Heisler, 1983; Holeton et al. 1983; Wright and Wood, 1985; Claiborne and Evans, 1988; Heisler, 1989). There is also evidence for a carrier-mediated transport of NH₄⁺via the electroneutral Na⁺/NH₄+ exchanger (Maetz and Garcia-Romeu, 1964; Kerstetter and Keeler, 1976; Payan, 1978; Wright and Wood, 1985), especially when the NH₃ gradient is directed inwards (Cameron and Heisler, 1983). In freshwater rainbow trout, acute exposure to alkaline water (pH9.5) strongly inhibited both ammonia excretion and Na⁺ influx (Wright and Wood, 1985). Water Na⁺ levels (Table 1) are much higher in Pyramid Lake water, which might facilitate Na⁺/NH₄⁺ exchange at high pH. Our results, however, indicate that the Na⁺/NH₄⁺ exchanger does not play a role in branchial ammonia excretion. Treatment with the Na⁺ uptake blocker amiloride or exposure to low-Na⁺ water had no effect on ammonia excretion (Table 3). In as much as our low-Na⁺ water still contained about 2.2mmol l⁻¹ of Na⁺, we cannot eliminate the possibility that a high-affinity Na⁺/NH₄⁺ exchanger may have persisted. However, in freshwater rainbow trout acclimated to 0.6mmol l⁻¹ Na⁺, the K_m for Na⁺ uptake was about 0.1mmol l⁻¹ (Goss and Wood, 1991). K_m generally scales with acclimation Na⁺ concentration, so the K_m for the present fish was probably about 10mmol l⁻¹, well above the concentration present in the low-Na⁺ water.

Another possible strategy to facilitate branchial ammonia excretion in alkaline water would be to enhance the passive loss of NH₃ across the gills by acidification of the water near the gill surface. Carbonic anhydrase on the gill surface (Rahim et al. 1988) catalyses the hydration of excreted molecular CO₂ to HCO₃⁻ and H⁺ and lowers the pH of the gill water boundary layer (Wright et al. 1986). Acetazolamide (1.6X10⁻³ mol l⁻¹), a carbonic anhydrase inhibitor, has been shown to reduce branchial ammonia excretion significantly in the freshwater rainbow trout (Wright et al. 1989). In contrast, acetazolamide had no effect on ammonia excretion rates in the present study (Table 3). Possibly, carbonic anhydrase is absent from the gill surface but, more likely, acidification of the gill water boundary layer is relatively small because of the high buffering capacity of the Pyramid Lake water.

If ammonia excretion is mostly due to the passive loss of NH₃ across the gills, then NH₃ levels in the plasma must surpass water NH₃ levels to maintain the blood-to-water diffusion gradients. Plasma levels depend on plasma pH and T_amm. Control blood pH (about 0.3pHunits higher) and plasma T_amm (about 100 µ,mol l⁻¹ higher) were both moderately elevated in the Lahontan cutthroat trout (Tables 4–6, Figs 3, 6) compared to the rainbow trout in neutral water (Wright and Wood, 1985; Wilkie and Wood, 1991), ensuring a positive gradient across the gills despite the high external pH of 9.4 (Figs 3, 4, 6). If ammonia excretion is dependent on this gradient, then acute changes in water pH should alter branchial ammonia excretion. Ammonia excretion was inhibited by 50% in fish acutely exposed to pH10 water (Fig. 2). Upon exposure to more acidic water (pH7.6), ammonia excretion rates increased threefold (Fig. 2). There was a sharp increase in the blood-to-water gradient when fish were initially exposed to pH7.6 water (0h, Fig. 4), coincident with a peak increase in ammonia excretion rates (0–1h, Fig. 4). Ammonia was effectively ‘washed out’ of the fish during the experimental period (0h, Fig. 3A), resulting in a marked reduction in both the gradient and ammonia excretion rate in the initial hour of the recovery period. The observation that J_amm remained significantly elevated over control rates throughout the 3h of exposure to pH7.6 water, despite a return of the gradient to control values, implies that mechanisms in addition to NH₃ diffusion were employed. One possibility is that a carrier-mediated NH₄⁺ mechanism was involved. Indeed, the increased availability of external H+ may have facilitated NH₄⁺ excretion via the putative H+/NH₄+ branchial transporter (Cameron, 1986). Clearly, further studies are required to identify the possible role of a Na⁺-independent NH₄⁺ transporter in these fish.

The NH₄⁺ electrochemical gradient was inversely correlated with ammonia excretion in fish exposed to pH7.6 water (Fig. 3). These results confirm previous findings that passive loss of NH₄⁺ is not a significant component of net ammonia excretion (Wright and Wood, 1985).

To investigate further the relationship between ammonia excretion and the gradient, fish were held in a closed system for 8h, resulting in a gradual build up of ammonia and urea levels in the water. There was a step-wise decrease in ammonia excretion in the first 6h of the experiment (Fig. 5), concomitant with a progressive increase in plasma total ammonia levels (Fig. 6) and a decrease in the gradient (Fig. 7). Surprisingly, ammonia excretion rates returned to ‘control’ rates (i.e. the 0–1 h value) after 6h, despite the fact that water total ammonia levels continued to rise. Moreover, the increase in ammonia excretion after 6h did not attenuate the accumulation of ammonia in the plasma. This may be due to changes in gill epithelial permeability or transport, in ammonia production rates, or to a redistribution of ammonia from intracellular stores to the extracellular compartment. The increase in ammonia excretion rates after 6h was associated with an increase in the gradient in some fish (Fig. 7). It may be that individual animals employed different mechanisms to restore ammonia excretion rates following several hours of depressed excretion while held in a closed system (Fig. 7). In some fish, a component of the increase in ammonia excretion after 6 h may have been due to a stimulation of carrier-mediated NH₄⁺ transport. Although ammonia excretion was independent of Na⁺ influx under conditions of normal plasma and water ammonia (Table 1), it may be that the Na⁺/NH₄⁺ exchange plays a role in net ammonia excretion when blood T_amm is elevated. In this regard, Wilkie et al. (1993) report that, during chronic exposure of Lahontan cutthroat trout to pH10 water, ammonia excretion was initially inhibited but later recovered and increased above control levels, at a time when plasma T_amm was persistently elevated. Furthermore, Yesaki and Iwama (1992) showed that, when rainbow trout were acutely transferred from neutral soft water to alkaline (pH10) hard water, ammonia excretion rates decreased, while plasma T_amm increased. The addition of amiloride to the alkaline hard water completely eliminated ammonia excretion over a 3h period, suggesting that, under these conditions, ammonia was excreted through a Na⁺-related mechanism (Yesaki and Iwama, 1992). There is also evidence in channel catfish (Ictalurus punctatus) that ammonia is excreted through an ion-exchange mechanism when water ammonia levels are elevated (Cameron, 1986). In Cameron’s study, ammonia excretion was independent of Na⁺, but appeared to be linked with H⁺ efflux. Thus, the increase in ammonia excretion in cutthroat trout following 6h in a ‘closed system’ may be explained, in part, by an increase in the transepithelial gradient and/or the induction of a carrier-mediated NH₄⁺ efflux mechanism.

To conclude, the Lahontan cutthroat trout do not employ any unusual physiological strategies for nitrogen excretion in alkaline water, but instead have modified processes typical of other teleosts. These modifications include lower rates of total ammonia excretion, a higher rate of renal ammonia excretion and a greater percentage of total nitrogen wastes excreted as urea compared to fish in neutral water. Ammonia excretion in Lahontan cutthroat trout is primarily dependent on the blood-to-water gradient. To overcome elevated water levels, plasma pH and T_amm are slightly elevated compared to those of fish in neutral water. Changes in ammonia excretion with changes in water pH and ammonia levels were associated, in part, with the transepithelial gradient, but carrier-mediated NH₄⁺ mechanisms may also play a role. Ammonia excretion is not dependent on Na⁺ influx or acidification of the gill water boundary layer under control conditions. These modifications are apparently not energetically costly, as Lahontan cutthroat trout grow to trophy size and support an active sport fishing industry.

We wish to thank Gary Wedemeyer of the US Fish and Wildlife Service and Paul Wagner, Lee Carlsen, Nancy Vucinich and Dan Mosely of the Pyramid Lake Fisheries for their enthusiastic cooperation and support during our stay. Maij Patrick, Russ Ellis and Jim McGeer all provided excellent technical assistance. We would also like to thank the Pyramid Lake Paiute Tribe for the invitation to study the unique fish of Pyramid Lake and for the opportunity to understand the successful fishery restoration program undertaken by the Paiute tribe since the 1970s. This research was supported by an NSERC International Collaborative Research Program awarded to G.K.I. and C.M.W. and NSERC operating grants to P.A.W., G.K.I. and C.M.W.