Physiological Responses of the Crayfish Pacifastacus Leniusculus to Environmental Hyperoxia : II. Role of the Antennal Gland in Acid–Base and Ion Regulation

In the preceding paper (Wheatly, 1989) it was shown that the extracellular acidosis occasioned by hyperoxia in the crayfish Pacifastacus leniusculus (Dana) was compensated by metabolic [HCO₃⁻] accumulation. This was accompanied by transbranchial exchange of acidic equivalents and electrolytes. In addition to the gills, decapods possess antennal glands (kidneys) which are primarily used for volume regulation in marine species since the urine produced is isosmotic (Riegel & Lockwood, 1961). In crayfish the antennal glands are unique in their ability to excrete a dilute urine (see reviews by Riegel, 1972; Mantel & Farmer, 1983); this is associated with the appearance of a morphologically distinct region of the distal tubule (Kamemoto & Tullis, 1972) where Na⁺ is actively reabsorbed by Na⁺/K⁺-ATPase (Peterson & Loizzi, 1974) followed by Cl⁻. Active solute reabsorption also occurs in the bladder (Kamemoto et al. 1962).

In higher vertebrates, urinary acid/base and electrolyte excretion are linked via exchange processes in the renal tubular cells (Hills, 1973). The specific activity of carbonic anhydrase (CA) in the crayfish antennal gland greatly exceeds that reported in marine species (Henry & Cameron, 1982; Wheatly & Henry, 1987), suggesting that it serves to provide counterions for Na⁺ and Cl⁻ reabsorption from an intracellular pool of CO₂. This implies that the antennal gland may be involved in acid-base regulation in the freshwater crayfish. Two previous studies have failed to demonstrate a comparable role in marine species (Cameron & Batterton, 1978; Wheatly, 1985).

The purpose of this study was therefore to correlate postfiltrational processing of electrolytes and acidic equivalents in the crayfish kidney during hyperoxia and recovery and compare any compensatory role with that of the gills. To do this, a non-invasive method for urine collection had to be developed.

Experiments were performed on adult intermoult crayfish Pacifastacus leniusculus leniusculus (Dana) (of either sex and mean mass 40·1 ± 3·9 g) maintained as previously outlined (Wheatly, 1989). The present paper describes findings from two experimental series. Series 3 used a preparation for direct collection of urine to measure effluxes of acidic equivalents and electrolytes for 48 h of control normoxia, 72 h of hyperoxia and 48 h of recovery. Series 4 used clearance of radiolabelled inulin to calculate urine flow rate (UFR) and subsequently determine electrolyte filtration and reabsorption rates under an identical experimental regime. Two of the eight animals used in series 4 were not fitted with a urinary collection device (UCD) to demonstrate that the procedure had no effect on inulin clearance.

Since crustaceans typically urinate intermittently and postfiltrational processing continues in the bladder, techniques for urine collection involving aspiration (Pritchard & Kerley, 1970) or internal cannulation of the ducts (Cameron & Batterton, 1978) were considered inappropriate. Initial attempts to cannulate the nephropores externally, as demonstrated in crabs (Holliday, 1977; Wheatly, 1985), were unsuccessful. Wherever this technique has previously been used in crayfish it has involved amputation of the chelae (Ono & Kamemoto, 1969; Tyler-Jones & Taylor, 1986). We wanted to avoid this procedure since our experiment required accurate determination of extracellular fluid volume (ECFV).

In preparing crayfish for urine collection, the flagellum of each second antenna was removed and the spines of the acumen, exopodite of the second antenna, and postantennal wing of the epistome were all filed down to prevent puncture of the collection bag. The crayfish was weighed and the chelae banded. A Qualatex grade 11 balloon, whose end had been cut to fit the contours of the anterior part of the protocephalic region, was then glued, with cyanoacrylate adhesive, ventrally to the epistomal ridge just posterior to the nephropore openings and dorsally just behind the eye orbits (Fig. 1). Before sealing the UCD at the sides, the balloon was flushed with nitrogen, which served the dual purpose of checking for leaks and, more importantly, preventing aeration of urine samples. Positioning of the balloon did not obstruct the mouthparts or occlude the exhalant branchial openings. Preliminary experiments indicated that complete recovery from surgical manipulation required 1 week. Patency and control urinary function were then ensured for 3–4 weeks (T. Toop & M. G. Wheatly, unpublished observations). Every 24 h mixed urine was removed from the balloon and UFR estimated from volume. A 26 gauge needle attached to a syringe was inserted through a resealing dental dam sampling port glued onto the surface of the balloon, which prevented subsequent leakage. The urine was analysed for acid-base parameters, electrolytes and inulin concentration.

In series 3, collected urine (u) was immediately assayed for pH, total carbon dioxide content and titratable acidity minus bicarbonate (TA – HCO₃⁻). The remainder was frozen and subsequently used to determine levels of ammonia, Na⁺, K⁺, Ca²⁺, Mg²⁺, Cl⁻, phosphate and sulphate.

Crayfish in series 4 were surgically prepared for postbranchial haemolymph removal 1 week prior to experimentation as outlined by Wheatly (1989). In this series, 48 h prior to the commencement of hyperoxia, the bath water was adjusted to a fixed volume (250 ml) with fresh water which was aerated and equilibrated to the experimental temperature. Each crayfish was then injected via the pericardial septum with 3 μCi of [³H]methoxyinulin (New England Nuclear; ECF marker repurified on Sephadex G-50 according to P. J. Walsh & C. M. Wood, personal communication) in 20 μl of crayfish saline. Every 24h for the 7 days of the experiment, the haemolymph, bath water and urine (in animals fitted with UCDs) were sampled and assayed for [³H]inulin radioactivity.

Urine pH (pHu) was determined on a 50- μ1 subsample using an IL 20985 liquid junction capillary electrode attached to a 213 blood gas analyser. was measured on a 40- μl subsample using the Capnicon (Cameron Instruments Inc.). Urine and bicarbonate concentration [HCO₃⁻ + CO₃²⁻] were calculated using the Henderson-Hasselbalch equation (see McMahon et al. 1978) and values for ⁠, and αCO₂ for a solution of comparable chlorinity (Harvey, 1974).

[TA-HCO₃⁻] was determined on 500 μl of fresh urine using the double titration procedure recommended by Hills (1973) and the IL pH microelectrode and blood gas analyser described above. The HC1 and NaOH titrants used were both 0·002 mol 1⁻¹ and the end point of the titration was taken as the corresponding mean haemolymph pH determined in series 1 (see Wheatly, 1989). Frozen samples were thawed and urine [ammonia] was determined on a 10-fold dilution using a micromodification of the phenolhypochlorite method of Solorzáno (1969). Urine acidic equivalent concentration was taken as the sum of the [TA — HCO₃⁻] and [ammonia] components (Hills, 1973).

Urine inorganic cation concentrations were determined using atomic absorption spectrophotometry (Perkin Elmer model 5000). Urine subsamples were diluted 1 in 1000 (Na⁺) or 1 in 30 (K⁺) with 0·1 % CsCl or 1 in 30 (Ca²⁺ and Mg²⁺) with 0·1 % LaCl₃.6H₂O to suppress interference in the air/acetylene flame. [Cl⁻] was determined on a 25- μl subsample of undiluted urine by coulometric titration (Radiometer CMT10). Total inorganic phosphate was measured on a twofold dilution of urine using a micromodification of the phosphomolybdic acid method of Atkinson et al. (1973). Inorganic sulphate was determined on a fivefold dilution of urine using the turbidometric method of Jackson & McCandless (1978).

For experimental series 4, samples of injection stock (20 μ1), haemolymph (20 ul), bath water (100 μl) and urine (20 μ;l) were diluted in 5 ml of fluor (ScintiVerse E; Fisher) and ³H radioactivity was measured on a scintillation counter (Beckman LS5801).

UFR was expressed in ml kg⁻¹ h⁻¹ and also in % BW day⁻¹ (where BW is body mass in grams).

Data are expressed throughout as mean ± S.E.M. (number of observations) and compared using Student’s two-tailed paired t-tests with 5 % as the fiducial limit.

Under control conditions urine was collected at rates of 4·30 ± 0·26 ml kg⁻¹ h⁻¹ in series 3 (Fig. 2). During the first 24 h of hyperoxia UFR virtually doubled; there was a slight accommodation with time although values remained 50% above control levels. UFR remained elevated during the initial 24h of recovery normoxia, thereafter returning to control levels.

Control urine concentrations of electrolytes were as follows (in mequivl⁻¹): Na⁺, 22·0 ±4·7; K⁺, 0·57 ±0·14; Ca²⁺, 1·55 ±0·17; Mg²⁺, 1·24 ±0·06; Cl⁻, 17·2 ±3·8; phosphate, 0·18 ±0·03; sulphate, 0·37 ±0·11; TA —HCO₃⁻, 0·23 ± 0·02; and ammonia, 1·74 ±0·11. Urinary effluxes of Na⁺ and Cl⁻ were comparable under control conditions (92·7 ±20·1 and 73·5 ± 9·5₍uequivkg⁻¹h⁻¹, respectively; Fig. 2) but responded differently to hyperoxia. Ė_Na tended to increase over the initial 48 h of hyperoxia, thereafter recovering, whereas Ė_Ci tended to become reduced after 36 h of hyperoxia. Both Ė_Na and Ė_C1 were significantly reduced initially on return to normoxia but subsequently recovered. Control urinary effluxes of the remaining haemolymph electrolytes were small by comparison (Fig. 3). Ė_K and Ė_Mg showed a similar response to hyperoxia to Ė_Na; excretion increased over the first 24 h of hyperoxia (by 44 % and 80 %, respectively) and decreased on recovery (by around 45 % ) before returning to control values. Eca exhibited a different trend; levels remained significantly elevated (by up to threefold) throughout hyperoxia and the first 24 h of recovery. Urinary phosphate excretion trebled during initial hyperoxic exposure, recovering partially after 48 h, somewhat resembling the profile exhibited by Ė_K and Ė_Mg. Renal sulphate efflux showed a similar trend but with a different time course; peak excretion rates (three times control) were approached between 24 and 48 h.

Significant changes were also observed in urinary acid-base parameters during hyperoxia (Fig. 4). Under control conditions the urine was more acid (pH 7·60) than the haemolymph (7·91; see Wheatly, 1989). During the first 24 h of hyperoxia the urine pH became significantly more acidic although control levels were return established within 48 h. Renal CO₂ efflux trebled initially, settling at a value which was double the control level for the duration of hyperoxia; control rates were rapidly re-established upon recovery. Around 90 % of the control net renal proton excretion was attributable to the nontitratable component, i.e. ammonia. Renal net proton efflux during hyperoxia and recovery mirrored ammonia efflux. Both doubled initially, progressively accommodated and recovered immediately upon return to normoxia. Renal [TA — HCO₃⁻] efflux remained at three times the control value for the duration of hyperoxia, once again recovering rapidly when mormoxia was reinstated.

In series 4 control crayfish, the rate constant (K) of inulin clearance was 0·71 ± 0·13 % h⁻¹. During hyperoxia K varied from 1·71 ± 0·26 % h⁻¹ in the initial sample to 1·82 ±0·29% h⁻¹ in the final sample. During initial recovery K was 1·12 ± 0·16 % h⁻¹, falling to control levels of 0·75 ±0·10% h⁻¹ by the 24–48 h sample. The ECFV of the crayfish in series 4 was calculated as 13·8 ± 1·5 ml which was 34·8 ± 2·2 % BW.

Control Ċ_IN was 4·5±0·6 mlkg⁻¹h⁻¹ (N=8) (10·7± 1·4% BWday⁻¹). Ċ_1N increased progressively throughout hyperoxia (Fig. 5), the rise becoming significant between 24 and 48 h and reaching peak values of 70% above control. Ċ_1N remained elevated for the first 24h of normoxic recovery. [IN]_U: [IN]_e was around 1·6 in control crayfish. This ratio tended to decrease during hyperoxia to levels of around 1·4 but returned to the control value upon recovery. To avoid aspirating bladder urine, the mean [IN]_U from the six cannulated crayfish was used to calculate UFR in the uncannulated animals. Calculated rates in the cannulated and uncannulated crayfish (Table 1) were similar, indicating that the presence of the urinary collection device did not interfere with inulin clearance rate. The combined control calculated UFR was 2·6 ±0·3 mlkg⁻¹h⁻¹ (N = 8) (6·9 ± 1·0 % BWday⁻¹) which agreed favourably with collected rates. UFR progressively increased throughout hyperoxia (Fig. 5) to peak values which were double the control rates and which remained elevated for 24 h of recovery. Therefore, similar trends in UFR were observed in both experimental series, although flow rates were generally lower in series 4 crayfish (Table 1).

Table 2 summarizes the postfiltrational reprocessing of various electrolytes by examining [X]_u: [X]_e. With the exception of ammonia, which is secreted, all other electrolytes had urine to haemolymph ratios below 1·6 (the value for inulin, see above) suggesting that they are reabsorbed. Under control conditions the ions most effectively reabsorbed were phosphate and sulphate (97 %) closely followed by Na⁺, Ca²⁺ and Cl⁻ at 95%. K⁺, Mg²⁺ and HCO₃⁻ were less effectively reabsorbed (85, 72 and 65 %, respectively).

Under control conditions Ṙ_Na and Ṙ_Cl each approached 1000 μequivkg⁻¹h⁻¹ (Fig. 6) which was an order of magnitude greater than the rate of reabsorption of any other ion. During hyperoxia, both progressively increased to peak values between 48 and 72 h, 60–80% above control levels which were re-established within 48 h of recovery. The remaining haemolymph cations exhibited similar trends except that Ṙ_K peaked within 48 h of hyperoxia and the increase in Ṙ_Mg was less pronounced (only 50%). During the initial 24h of recovery, reabsorption of all these ions remained high; recovery again occurred within 48 h. The time course of Ṙphosphate paralleled that of Ṙ_Na except that peak values were 150 % above the control level. Ṙ_sulphate exhibited a linear increase with time in hyperoxia, becoming significantly elevated 24 h earlier than Ṙphosphate but with peak values again increased by 150%. However, control reabsorption rates were re-established more rapidly. was significantly elevated throughout the hyperoxic period with peak reabsorption rates of four times control values. Reabsorption rates remained elevated for the initial 24 h of recovery.

Control ammonia secretion rate was low compared with rates of reabsorption (Fig. 7). Ṡ_ammonia increased progressively during hyperoxia, becoming significantly elevated (50 % above control) in the final collection period. This trend was rapidly and immediately reversed upon recovery when the rate of secretion dropped significantly below control levels and subsequently recovered.

Close agreement between UFR calculated from Ċ_IN and from collected flow (Table 1) in series 4 crayfish confirms that the UCD developed in this study was satisfactory. Agreement between Ċ_IN values in cannulated and uncannulated crayfish suggests that the UCD did not affect UFR. The combined calculated and collected value for UFR (7·1 ± 1·0% BWday⁻¹) agrees with flow rates determined by direct external cannulation in another crayfish study (Tyler-Jones & Taylor, 1986). Earlier studies estimating UFR by weight gain following nephro-pore blockage (e.g. Maluf, 1941) have consistently yielded lower values, confirming that increased back pressure reduces filtration. Flow rates calculated in previous investigations are not strictly comparable with the present study if [IN]_u:[IN]_e was determined on aspirated bladder urine. In marine species, water is reabsorbed in the bladder so that aspirated urine typically has a lower [IN]_U: [IN]_e value than voided urine (see Wheatly, 1985) and this would tend to overinflate UFR. The reverse situation occurs in the crayfish. The [IN]_U: [IN]_e in voided urine (1·6, present study and Tyler-Jones & Taylor, 1986) was much lower than values previously reported for aspirated urine (2·5, Riegel, 1961; Pritchard & Kerley, 1970). We are presently unable to explain this discrepancy. The fact that [IN]_u:[IN]_e is above unity indicates net reabsorption of water in the antennal gland despite selective pressure to eliminate a water load; presumably this is an unavoidable consequence of active solute reabsorption. There was, however, good agreement between Ċ_IN determined in this study and values reported by Tyler-Jones & Taylor (1986) and Riegel (1961). Pritchard & Kerley (1970), however, reported lower values when monitoring the appearance of radioinulin in the bath water rather than disappearance from the haemolymph.

The increase in UFR (Figs 2,5; Table 1) was similar though more pronounced than that reported in a comparable hyperoxic study in trout (Wheatly et al. 1984). Two factors contributed to this diuresis: a reduction in net water reabsorption ([IN]_u:[IN]_e decreased) and an increased clearance (Ċ_IN, Fig. 5). The latter is generally attributed to an increase in haemolymph volume or pressure. In a later part of this study we found no evidence for an increase in circulating volume (M. G. Wheatly, R. Morrison, T. Toop & L. C. Yow, in preparation) and we did not measure intracardiac pressure (equivalent to antennary artery filtration pressure) in this study. However, in a parallel study on rock crabs (M. G. Wheatly, in preparation) hyperoxia was accompanied by a maintained bradycardia which has been correlated with increased ventricular pressure in decapods (Burggren et al. 1985; deFur et al. 1985). An alternative explanation for the increase in UFR is that the permeability of the filtration membrane is altered by a diuretic hormone. Norfolk & Craik (1980) similarly concluded that a diuretic factor may be involved in increasing UFR in crabs in dilute sea water, since the increase could not be explained by changes in arterial pressure.

Urine electrolyte concentrations agreed with previous determinations in voided crayfish urine (Tyler-Jones & Taylor, 1986); lower levels have been reported in aspirated urine (e.g. Pritchard & Kerley, 1970) consistent with reduced flow rates. One common feature is that [Cl⁻]_u, is consistently lower than [Na⁺]_u, suggesting that minor anions contribute more to net negativity than minor cations do to net positivity. However, the resulting anion gap (i.e. Na⁺ + K⁺ + Mg²⁺ + Ca²⁺ − Cl⁻ − PO₄²⁻ − SO₄²⁻ − HCO₃⁻) is relatively small, indicating that the major urinary electrolytes were identified in this study.

The antennal gland appears to act in concert with the gills (Wheatly, 1989) in the control whole-body regulation of certain ions such as Na⁺ and Cl⁻ which undergo net losses of around 100 μequiv kg⁻¹ h⁻¹ at each epithelium. Renal Ca²⁺ excretion is small compared to net branchial loss. In addition, renal excretion of other ions (e.g. K⁺ and Mg²⁺) is offset by net branchial uptake. The dynamics of whole-body ion regulation appear to alter during experimental hyperoxia. For example, renal excretion of Na⁺ and Cl⁻ countered branchial net fluxes, and Mg²⁺ was handled similarly, constituting a reversal of the control characteristics. Calcium was exceptional in that the kidney predominated over the gills in terms of whole-body efflux during the experiment.

In contrast to marine species (Cameron & Batterton, 1978; Wheatly, 1985), control crayfish produce a urine that is acid with respect to the haemolymph. Our measurements agree with an isolated measurement from Maluf’s historic article (1941) which reports a pH for bladder urine of 7·5. Moreover, the components of urine acidity differ in magnitude and in some cases vector. In a marine crab, such as Cancer (Wheatly, 1985), the alkaline pH is primarily attributable to a titratable alkalinity efflux (5·0 μequivkg⁻¹h⁻¹) which is negligibly countered by ammonia efflux (0·2 μequiv kg⁻¹h⁻¹). In crayfish, urine acidity is primarily determined by ammonia efflux combined with a small excretion of titratable protons (Fig. 4). Although the magnitude of renal ammonia efflux differs between these two species, whole-animal excretion rates are comparable (Wheatly, 1989, in preparation). Control freshwater fish also produce an acid urine (Wheatly et al. 1984). Calculating the ratio of nontitratable to titratable effluxes emphasizes the relative importance of ammonia in the renal excretion of acid in freshwater species. This value is around 7·39 in crayfish and 1·5-6·5 in trout; both are considerably in excess of the typical mammalian value (1–2·5).

Our data suggest that the crayfish antennal gland, by increasingly acidifying the urine (Fig. 4), assisted the gills in correcting the extracellular hyperoxic acidosis. The magnitude of the total renal response was only 500 μequivkg⁻¹ which is approximately 10% of the net branchial H⁺ loss (Wheatly, 1989), agreeing with other studies on the relative role of the gills and kidney in freshwater species (Heisler, 1984). H⁺ was excreted predominantly as ammonia although the titratable component of urine acidity also increased.

Although the two primary exchange epithelia exhibited comparable net electrolyte fluxes, the antennal gland had a greater capacity for ion reabsorption. Control values for Ṙ_Na and Ṙ_Cl (Fig. 6) significantly exceeded corresponding values for branchial unidirectional influx (⁠ and ⁠, Wheatly, 1989). If the ion uptake mechanisms are equally energy-dependent then, as Kirschner (1967) suggests, the costliest part of hyperosmotic regulation should be operating the kidneys. The difference in ion-transporting capability is further amplified if one considers the relative surface area of the two exchange organs. The branchial surface area of a 35-g crayfish can be estimated as 140 cm² (McMahon & Wilkens, 1983). Based on Maluf’s original observations (1941) the combined exchange area of the nephric tubules is approximately 3·76cm². The exchange area ratio for gills: kidney is therefore 40:1. If the antennal gland achieves three times the net ion reabsorption with one-fortieth of the surface area, then the specific activity of transport enzymes per unit area should be 120 times greater. Wheatly & Henry (1987) confirmed a threefold discrepancy between antennal-gland- and gill-specific activities of Na⁺/K⁺-ATPase and carbonic anhydrase expressed per gram of tissue homogenate protein. The ratio of ammonia loss at the gills to that at the kidney (65:2·3) may also reflect relative exchange areas (all remaining diffusional characteristics being equal). The nephric tubule is less elongated in marine species which may explain why their renal ammonia effluxes are lower than that of the crayfish. Electrolyte reabsorption and secretion rates showed a more pronounced response to hyperoxia than did excretion rates. Just as for net flux rates, the antennal gland and gills (Wheatly, 1989) exhibited opposing trends with respect to unidirectional influx rates of Na⁺ and Cl⁻ during hyperoxia. In other cases (e.g. ammonia) epithelia worked in concert although an earlier response time was generally seen at the gills.

Together with inulin clearance (Fig. 5) the present data would suggest that glomerular filtration and tubular reabsorption rate are balanced in an attempt to regulate circulating electrolyte levels. Fluid balance, however, did not appear to be regulated by feedback control, as is typical in the mammalian nephron, although UFR partially recovered in series 3 crayfish (Fig. 2). In fact the [IN]_u:[IN]_e ratio suggested that fluid reabsorption decreased. The primary stimulus for tubular fluid reabsorption in mammals is increased plasma colloid osmotic pressure (COP). This stimulus may be less effective in crustaceans since they excrete protein in the urine (Kirschner & Wagner, 1965).

The renal response to respiratory acidosis in the crayfish was urinary acidification and net HCO₃⁻ reabsorption which assisted in alkalinizing the blood.

Although this is the same overall response seen in mammals (Marsh, 1983), there appear to be some differences between the mechanisms involved. In mammals, the two buffers involved in tubular acid secretion (which ultimately form new HCO₃⁻ in the blood) are phosphate (titratable acidity) and ammonia (non-titratable acidity). In the tubular lumen, the dibasic phosphate (HPO₄²⁻) accepts a proton, forming monobasic H₂PO₄⁻ which is then excreted. When total phosphate excretion in the present study was resolved into dibasic and monobasic components (Fig. 8) by means of the Henderson-Hasselbalch equation (at urine pH and assuming a of 6·8) it transpired that H₂PO₄⁻ only accounted for 10% of the [TA - HCO₃⁻] efflux. It would therefore constitute a minute fraction of the TA flux given that urinary CO₂ is very high. This strongly suggests that other urinary buffers, perhaps including urinary proteins (Kirschner & Wagner, 1965; M. G. Wheatly, unpublished observations), contribute to titratable acidity.

The urinary NH₃/NH₄⁺ buffer pair appears to play the more important role in crayfish during hyperoxia. Renal ammonia excretion (Fig. 4) was precipitated by the fall in urine pH which increased diffusional loss of NH₃ by increasing the gradient. After protonation, NH₄⁺ is typically excreted as a neutral salt. Since renal Cl⁻ efflux remained constant (Fig. 2) it is unlikely that NH₄⁺ was excreted as NH₄C1. However, sulphate excretion (Fig. 3) paralleled ammonia excretion obeying a 1:2 stoichiometry, making (NH₄)₂SO₄ excretion a distinct possibility. Electrolyte handling by cells in the antennal gland has been confirmed by subsequent intracellular determinations (M. G. Wheatly & E. C. Vevera, in preparation). Cellular ammonia and sulphate levels were elevated whereas phosphate level remained unchanged.

Presumably proton secretion into the lumen from the renal tubule cell is analogous in crustaceans and vertebrates, occurring via apical carrier-mediated electroneutral ion exchange, i.e. H⁺/Na⁺. Basolateral HCO₃⁻ reabsorption will include newly formed HCO₃⁻ and filtered HCO₃⁻. Both will acidify the urine but the latter will not change circulating HCO₃⁻ levels. Although HCO₃⁻ reabsorption (Fig. 6) adapts to the filtered load, any which escapes reabsorption will raise pH, thereby increasing the driving force for proton secretion. A reduction in ECFV which occurs initially during hyperoxia (M. G. Wheatly, R. Morrison, T. Toop & L. C. Yow, in preparation) is also a powerful stimulus for proton secretion, again acting via the Na⁺/H⁺ counter-transporter mechanism. However, the principal stimulus for renal acidification is ⁠, which increased in the renal cells (M. G. Wheatly, R. Morrison, T. Toop & L. C. Yow, in preparation) and might increase the driving force for proton secretion by lowering cell pH.

From the studies which exist, monobasic phosphate would appear to be the major urinary buffer in fish, as in mammals. Trout showed a long-lived urinary acidification during hyperoxia (Wheatly et al. 1984); however, the net acid efflux reflected changes in titratable acidity, not ammonia levels. Furthermore, the TA efflux was stoichiometrically equivalent to H₂PO₄⁻efflux, which was increased via a switch from net phosphate reabsorption to secretion.

We thank Dr Chris Wood, McMaster University, Canada, for his helpful comments on the manuscript, and Regan Meeker for assisting us with many of the assays. The following, at the University of Florida, are gratefully acknowledged for the loan of experimental equipment: Dr J. J. Delfino (Environmental Engineering), Dr D. H. Evans and Dr F. G. Nordlie (Zoology) and Dr Thomas (VA Hospital). Grace Kiltie prepared the manuscript and Daryl Harrison the figures. This study was funded by NSF grant no. DCB 84-15373 to MGW.