Determination of NH+4/NH3 fluxes across apical membrane of macula densa cells: a quantitative analysis

M. Anuar Laamarti and Jean-Yves Lapointe

Groupe de Recherche en Transport Membranaire, Université de Montréal, Montréal, Quebec, Canada H3C 3J7

    ABSTRACT
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Abstract
Introduction
Materials & Methods
Results
Discussion
Appendix
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Luminal addition of 20 mM NH+4 produced a rapid acidification of rabbit macula densa (MD) cells from 7.50 ± 0.06 to 6.91 ± 0.05 at an initial rate of 0.071 ± 0.008 pH unit/s. In the luminal presence of 5 µM bumetanide, 5 mM Ba2+ or both, the acidification rate was reduced by 57%, 35% and 93% of control levels. In contrast, intracellular pH (pHi) recovery after removing luminal NH+4 was unaffected by bumetanide and Ba2+ but was sensitive to 1 mM luminal amiloride (71% inhibition). The bumetanide-sensitive acidification rate represents most certainly the NH+4 flux mediated by the apical Na+:K+ (NH+4):2Cl- cotransporter, but the Ba2+-sensitive portion does not seem to be associated with the apical K+ channels previously characterized by us. The effects of NH+4 entry across the apical membrane were simulated using a simple model involving five adjustable parameters: apical and basolateral permeabilities for NH+4 and NH3 and a parameter describing a pH-regulating mechanism. The model shows that the apical membrane of MD cells is much more permeable to NH3 than it is to NH+4 and, under control conditions, the apical NH+4 flux appears surprisingly high (11-20 mM/s) and challenges the notion that MD cells present a low intensity of ionic transport.

ammonium permeability; sodium-potassium-chloride cotransporter; potassium channels; bumetanide; verapamil

    INTRODUCTION
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Abstract
Introduction
Materials & Methods
Results
Discussion
Appendix
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THE MACULA DENSA (MD) is a plaque of epithelial cells located at the distal end of the thick ascending limb (TAL) that is thought to function as a sensor device detecting increases in luminal NaCl concentrations and initiating the signals involved in the control of renin secretion and in the initiation of the tubuloglomerular feedback (24). Because of their small number (~30 cells/plaque), MD cells could not be directly studied using conventional methods, and their transport properties remained largely unknown until 1985, when it was demonstrated that MD cells could be visualized during microperfusion experiments of isolated TAL dissected with their attached glomerulus (15). Their transport properties could then be studied using fluorescent probes (7, 19, 20, 21), conventional electrophysiology (4, 17, 18, 23), and patch-clamp techniques (9, 22). The transport model that emerged from these studies consists of Na+:K+:2Cl- cotransporters, Na+/H+ exchangers, and K+ channels on the apical membrane and a major Cl- conductance, a K+ conductance, Na+/Ca2+ exchangers, and Na+-K+-adenosinetriphosphatases (Na+-K+-ATPases) on the basolateral membrane. Even though the proposed model for MD cells is qualitatively similar to the cortical TAL (CTAL) model, differences were reported on the basis of the properties of the apical K+ channels observed at the single channel level (9) and on the Na+:K+:2Cl- isoform detected in MD cells (10). Also, CTAL cells were presumed to be much more active than MD cells in terms of absolute transport rates, as the density of basolateral Na+-K+-ATPase was estimated to be 40 times smaller in MD cells vs. CTAL cells (reported per unit of cell volume) (Ref. 25; see also Refs. 3 and 11). So far, the level of membrane or transepithelial transport mediated by MD cells has never been measured. More information on MD cells properties, including membrane transport properties, is required to understand the way these cells play their crucial role in the kidney.

Recently, we have presented a method to detect the direction of ionic flux mediated by the apical Na+:K+:2Cl- cotransporter using intracellular pH (pHi) measurements (19). The method gave interesting results but was quite indirect as pHi was shown to be linked to the apical Na+:K+:2Cl- flux through the modulation of intracellular Na+ concentration ([Na+]i) and the activity of the Na+/H+ exchanger. In the present study, we apply to MD cells a method previously used for medullary TAL (MTAL) that is based on the fact that NH+4 can substitute for K+ in several membrane transporters and channels including the Na+:K+:2Cl- cotransporter (14) and directly affect pHi by dissociating into NH3 + H+. It will be shown that luminal addition of NH+4 produces a rapid cellular acidification that can be almost completely inhibited by bumetanide and Ba2+. A simple model is presented for the quantitative interpretation of these results. The rate of NH+4 entry across the apical membrane was found to be surprisingly high, which suggests that, even if MD cells have a low density of basolateral Na+-K+-ATPase, their apical membrane presents a large ionic permeability.

    MATERIALS AND METHODS
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Abstract
Introduction
Materials & Methods
Results
Discussion
Appendix
References

Microperfusion. Microperfusion of rabbit CTALs dissected with their attached glomeruli was performed as described in previous reports from this laboratory (4, 17, 18, 19). The distal end of the tubule was left open to the bath, and a holding pipette was placed over the glomerulus to position the preparation in such a way that the MD plaque could be clearly visualized using a ×40 objective. Tubules were bathed with a bicarbonate-free solution containing (in mM) 146 NaCl, 5 potassium gluconate, 1 MgCl2, 1 CaCl2, 5 glucose, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), and 7.2 tris(hydroxymethyl)aminomethane (Tris). Luminal solutions were identical to the bathing solution with the exception that the luminal NaCl concentration was maintained at 25 mM by isosmotically replacing Na+ with N-methyl-D-glucamine (NMDG) and Cl- with cyclamate. Addition of luminal NH+4 (20 mM) and or Ba2+ (5 mM) was accomplished by replacement of NMDG-cyclamate at constant luminal [Na+] and [Cl-]. In some experiments, 5 µM bumetanide, 1 mM amiloride, 1 µM ionomycin, or 0.1 mM verapamil (all compounds from Sigma Chemical, St. Louis, MO) was added to the perfusion solution from concentrated solutions in ethanol (final ethanol concentration in perfusates was <0.2%). All solutions were adjusted to a pH of 7.4, and all experiments were performed at 39°C.

Fluorescence measurements. pHi was measured using the fluorescent probe 2',7'-bis(carboxyethyl)-5(6)-carboxyfluorescein (BCECF) as previously described (7, 19). In brief, after a CTAL was cannulated and perfused on the stage of an inverted microscope, background fluorescence was measured over a window positioned over the MD plaque, and 5 µM of the acetoxymethyl ester form of BCECF (BCECF-AM) was added to the luminal perfusate. Intracellular dye was excited alternately at 500 and 450 nm wavelengths (Spex model CM-III; Spex Industries, Edison, NJ), and fluorescence emission was monitored at 530 nm using a photomultiplier tube and a band-pass filter. BCECF-AM was not removed from the lumen until the fluorescence measured for both excitation wavelengths had increased by a least one order of magnitude with respect to background fluorescence.

BCECF fluorescence was calibrated using the high-K+ nigericin technique (26). At different periods during our study, a total of seven tubules were perfused and bathed with identical solutions containing (in mM) 120 KCl, 1 MgCl2, 2 NaH2PO4, 25 NaCl, 0.006 nigericin, and a mixture of Tris and HEPES (25 mM total) giving pH values of 6.4, 6.8, 7.2, 7.6, and 8.0. Calibration curves were obtained, and one can see that the fluorescence ratios converge to the same value in acidic conditions (6.4-6.8), whereas all the ratios were distributed within ± 7% of the mean at a pHi of 7.6. As a complete calibration could not be performed in each experiment, one or two calibration points (including pHi = 7.6) were obtained at the end of each experiment, and the calibration curve which best satisfied these points was used to scale the measured fluorescence ratios.

Buffering capacity. The buffering capacity of the cell (beta i) was determined from the following equation using the measured change in pHi when 10 mM of trimethylamine (TMA) was removed from the luminal solution.
&bgr;<SUB>i</SUB> = &Dgr;H/&Dgr;pH
where Delta H is the concentration of proton released when TMA is removed from the luminal solution as calculated from pHi and a pKa of 9.83. This measurement was performed by adjusting the luminal and basolateral solution pH to three different values (6.4, 7.4, and 8.0) to estimate beta i at a variety of pHi.

Acidification rates and flux units. Initial rates of pHi change (dpHi/dt) were calculated from a fit (FigP, version 6.0; Biosoft, Milltown, NJ) of the recorded pHi to an exponential relaxation curve for the initial 20-30 s following a change in luminal solution. On a few occasions where changes in pHi were clearly not exponential during this period, a linear regression was fitted to the initial change in pHi. Acidification rates can be transformed into proton production rates in units of millimolar per second by simply multiplying dpHi/dt by beta i. Similarly, ionic fluxes can be expressed in the same convenient units (mM/s), which directly indicates the rate at which an intracellular concentration would change following a given transmembrane flux. Care should be taken, however, in comparing ionic fluxes among different cell types, as cellular volume may vary quite significantly from one cell type to another.

Statistics. Statistical significance of a difference between two average results was tested using Student's t-test for paired sample. P < 0.05 was considered significant.

    RESULTS
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Buffering capacity. Intrinsic buffering power was obtained as described above using 10 mM TMA. As displayed in Fig. 1, one can see that beta i increases very significantly from 16 to 141 mM/pH unit as pHi decreases from 8.1 to 6.7. This can be directly appreciated in the inset of Fig. 1; with a pKa of 9.83 for TMA, the quantity of proton released upon removal of TMA is slightly larger (by ~10%) at pHi 6.4 than at pHi 7.8; however, the acidification induced by TMA removal was much smaller at pHi 6.4 than at 7.8. For each experiment, the conversion between dpHi/dt and proton production-rate was calculated using interpolated beta i estimates as depicted in Fig. 1.


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Fig. 1.   Buffering capacity of macula densa (MD) cells. MD cells intracellular pH (pHi) was varied by use of different extracellular pH (from 6.4 to 8.0), and buffering capacity was determined by measuring the change in pHi immediately following removal of 10 mM trimethylamine (TMA) from the luminal solution. Data points are means ± SE, and for each, the number shown in parentheses is the number of MD plaques studied.

Effects of luminal ammonium addition on pHi. In the absence of ammonium, the steady-state pHi was 7.50 ± 0.06 (n = 8) when the CTAL lumen was perfused with a low NaCl concentration (25 mM). Addition of 20 mM NH+4 to the luminal perfusate caused occasionally a small increase of pHi [average of +0.02 ± 0.009 pH unit, not significant (NS)], which was followed by a rapid cell acidification (Fig. 2A) at an initial rate given in Table 1. A final pHi of 6.91 ± 0.05 was usually reached within 30 s. Removal of luminal NH+4 caused an additional but transient acidification by an average of 0.058 ± 0.015 pH unit (n = 8) followed by a cellular alkalinization at a rate corresponding to about one-third of the absolute value of the initial NH+4-induced acidification (see Table 1) and a stable pHi of 7.53 ± 0.06 was reached within 90 s.


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Fig. 2.   Effects of adding 20 mM luminal NH+4 on MD pHi. The experiment was done for the same MD plaque in control conditions (A) or in presence of 5 µM bumetanide (B), 5 mM Ba2+ (C), or both (D).

                              
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Table 1.   Initial pHi and acidification rate induced by luminal addition/removal of 20 mM NH+4

Effects of NH4 in presence of bumetanide and Ba2+. To determine whether apical Na+:K+:2Cl- cotransporters are involved in NH+4 transport, the effect of adding 20 mM NH+4 to the lumen was evaluated in the presence of 5 µM bumetanide. A typical tracing of the effect of bumetanide is shown in Fig. 2B. As summarized in Table 1, bumetanide produced a very significant effect on NH+4-induced acidification (P < 0.002, n = 5), which was reduced by 57%, in the presence of the inhibitor. Time control experiments were performed to check the reproducibility of the NH+4-induced acidification. In a series of four different experiments in which NH+4 was added and removed twice at 5- to 10-min intervals, the second NH+4-induced acidification was not significantly different from the initial acidification (the average change in the initial acidification rate was +5 ± 9%; P = 0.50, NS).

Thus apical Na+:K+:2Cl- cotransporters are likely to be involved in the NH+4 entry mechanism, but the incomplete effect of bumetanide suggests the presence of a second pathway. Since K+ channels are present at the apical membrane of MD cells (9), and since Ba2+ is an effective inhibitor of several K+ channels and was shown to significantly inhibit NH+4-induced acidification in rat MTAL cells (8, 12, 13, 27), the effect of 5 mM Ba2+ was tested on MD cells. In eight tubules, addition of 5 mM Ba2+ to the luminal medium reduced the NH+4-induced acidification by 35% compared with control values (Fig. 2C; Table 1). Interestingly, the effects of bumetanide and Ba2+ appeared additive, as the simultaneous presence of 5 µM bumetanide and 5 mM Ba2+ inhibited the NH+4-induced cell acidification nearly completely, reducing the initial acidification rate by 93% (Table 1; Fig. 2D).

The results presented above are similar to the results reported for MTAL (8, 12, 13, 27) and were interpreted in the past as evidence that NH+4 is indeed entering through apical Na+:K+:2Cl- cotransporters and Ba2+-sensitive K+ channels. On the other hand, it was shown for rat TAL that their apical K+ channels do not conduct NH+4 (5, 6), and it was recently suggested that, in the case of suspensions of rat MTAL, NH+4 was able to replace the proton in a K+/H+ exchanger that was sensitive to Ba2+ and verapamil (1). As we have recently reported in a series of patch-clamp experiments that the apical K+ channels of MD cells could be inhibited by a rise in intracellular Ca2+ induced by application of 1 µM ionomycin (9), we tested the effect of ionomycin on the Ba2+-sensitive NH+4 transport through the apical membrane of MD cells. The results are shown in Fig. 3. In this series of experiments, bumetanide and Ba2+ reduced the NH+4-induced acidification rate to 10% of its control value (n = 12). Clearly, 1 µM ionomycin could not mimic the effect of Ba2+ on NH+4-induced acidification, as an acidification rate corresponding to 79% of the control acidification rate could be recorded (see Fig. 3 for an example), which suggests that the Ba2+-sensitive NH+4 influx is not mediated by the K+ channels that we previously observed in patch-clamp experiments (9). In the following series of experiments, the effect of 0.1 mM verapamil was tested in the presence of 5 µM bumetanide and in the presence of 5 mM Ba2+ to see its effect through the putative K+/H+ (NH+4) exchanger on each of the two components of the NH+4-induced acidification. In six experiments, the NH+4-induced acidification rate was shown to be 0.073 ± 0.010 pH unit/s in the presence of bumetanide and verapamil, a value not significantly different from the value of 0.090 ± 0.013 pH unit/s obtained in the presence of bumetanide alone (P = 0.24). In the presence of Ba2+, verapamil also failed to significantly affect the acidification rate (0.076 ± 0.025 pH unit/s before and 0.070 ± 0.024 pH unit/s after the addition of verapamil, P = 0.14, n = 5).


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Fig. 3.   Comparison between effect of ionomycin and effect of Ba2+ on NH+4-induced acidification. NH+4, 20 mM, was added to luminal perfusate in control conditions, in presence of 5 µM bumetanide + 1 µM ionomycin, or 5 µM bumetanide + 5 mM Ba2+.

pHi recovery after NH+4 removal. To determine whether NH+4 could exit MD cells via the pathways involved in the entry process, the pHi recovery during NH+4 washout was evaluated in the presence of bumetanide and Ba2+ (Table 1). Under control conditions, alkalinization rate during the recovery period was not changed by the presence of bumetanide and Ba2+ (Fig. 4A; Table 1). If amiloride was present in the lumen, then the alkalinization rate was reduced to 29% of the control value indicating a dominant role for the apical Na+/H+ exchanger in pHi recovery following NH+4 removal (Fig. 4B; Table 1).


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Fig. 4.   pHi recovery during luminal NH+4 wash out in MD cells. A: for a single MD plaque, there was an absence of any significant effect of 5 µM bumetanide + 5 mM Ba2+ in the lumen on the rate of pH change during the recovery period. B: effect of 1 mM luminal amiloride on the alkalinization rate following luminal NH+4 removal. At the vertical arrow, amiloride was removed from the perfusate.

    DISCUSSION
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Comparison between NH+4-induced acidification in MD vs. TAL cells. The results presented above indicate that, qualitatively, MD cells behave like mouse (12, 13) and rat (1, 27) MTAL cells when their apical membrane is exposed to NH+4. In the cases where microperfusion was used, a dominant acidification was systematically recorded after luminal NH+4 addition, which could be blocked almost completely by furosemide and Ba2+ (12, 13, 27). For MTAL suspensions (1, 12, 13), similar acidifications were observed after bilateral NH+4 addition, but ouabain was needed in addition to Ba2+ and furosemide to block the acidification completely. Our average acidification rate under control conditions (0.071 pH unit/s) compares with published determinations in MTAL: 0.047 pH unit/s for microperfused mice MTAL (13), 0.053 pH unit/s for rat MTAL suspension (1), and 0.185 pH unit/s for microperfused rat MTAL (27).

Conversion of acidification rates into net proton production (JH) requires an estimate of the buffering capacity for each cell type. We have found a steep dependence of beta i as a function of pHi in MD cells that is obvious from the raw data (see inset of Fig. 1). The average beta i of MD cells at a pHi of 7.5 is 70 mM/pH unit. In mouse MTAL cells, beta i was also shown to increase with cellular acidification and averages 29.7 mM/pH unit in the pHi range of 7.0 to 7.6 (13). Higher beta i values were estimated for rat MTAL (85 mM/pH unit, unpublished data cited in Ref. 27) whereas in rabbit proximal tubules (16), beta i was estimated to 42.8 mM/pH unit (84.6 mM/pH unit in the presence of CO2/HCO<SUP>−</SUP><SUB>3</SUB>). In consequence, net proton production rates after adding luminal NH+4 (dpHi/dt × beta i) were 5.0 mM/s for MD cells in control conditions, 1.4 mM/s for mice MTAL (13), and 15.7 mM/s for rat MTAL (27), as estimated from their unpublished value for beta i. One can see that the net proton production rate may vary considerably depending on the species and on the buffering capacity estimated in each case. Nevertheless, proton production rate in MD cells induced by NH+4 addition is clearly not 40 times smaller than the corresponding values for MTALs. This is in contrast with the low intensity of ionic transport expected for MD cells, given the fact that the MD Na+-K+-ATPase activity was estimated to be 1/40 of the TAL activity (expressed per unit of cell volume as is also the case of proton production rates; see Ref. 25; see also Ref. 3). However, the interesting parameters to estimate are the NH+4 and NH3 fluxes (JNH4 and JNH3), and the following model will help us in going from JH to JNH4 and JNH3.

Transport model for simulating NH+4/NH3 fluxes in epithelial cells. It has been previously assumed that a powerful intracellular acidification following addition of NH+4 indicated the presence of an NH+4 permeability (PNH4) much larger than the NH3 permeability (PNH3) (12, 27) and that the proton production rate (dpHi/dt × beta i) could be directly assimilated to the net flux of NH+4 (JNH4) (13). However, as recognized by Watts and Good (27), with a pKa of 9.0 for the NH+4 dissociation reaction and a pHi of 7.4, only 1/40 (~10-1.6) of NH+4 ions entering the cell should dissociate to NH3 + H+. In the case of mouse MTAL, the proton production rate would then correspond to a JNH4 of 56 mM/s, a clearly unacceptable value that is inconsistent with the fact that pHi and presumably intracellular NH+4 concentration require up to 30 s to reach a steady-state level. The model briefly presented here and in more detail in the APPENDIX will show that both of these assumptions (PNH4 > PNH3 and JNH4 = proton production rate) are wrong in the case of MD cells and most likely in the case of MTAL cells as well.

We are considering a simple model that takes into account a pHi regulation system together with NH3 and NH+4 fluxes across apical and basolateral membrane. Permeabilities are defined as the coefficient (in s-1) by which a cis concentration (mM) has to be multiplied to obtain a unidirectional flux (mM/s) in the trans direction. In the case of a simple diffusion of neutral substrate (as for NH3, for example) permeability coefficients for influx are naturally set equal to the corresponding coefficient for efflux across a given membrane. In the case of NH+4, however, provisions are made to allow different permeability coefficients for influx and efflux in such a way that membrane potential and/or cotransported substrates have the possibility to generate asymmetrical fluxes producing intracellular NH+4 accumulation. The Na+/H+ exchanger that has been recently identified in the apical membrane of MD cells (7) was arbitrarily modeled in such a way that the proton efflux was made proportional to the value of 7.5 - pHi, which roughly mimics the general function of an Na+/H+ exchanger in the presence of a constant Na+ gradient (2). In the absence of basolateral NH3/NH+4, the five parameters to adjust are as follows: the apical and basolateral permeabilities for NH3 (PaNH3 and PblNH3, respectively), the apical NH+4 permeability for influx (PaiNH4), the sum of basolateral and apical NH+4 permeability for efflux (PeNH4) and the pHi sensitivity (SH) of the pHi-regulating mechanism [proton efflux being given as SH × (7.5 - pHi)]. In this simple model, the dissociation of NH+4 was assumed to be sufficiently fast to continuously keep intracellular NH+4 close to the equilibrium with intracellular NH3 and H+. Finally, the measured buffering power was represented as a linear function of pHi that corresponds to the measured values between a pHi of 6.7 and 8.1 (Fig. 1) (see APPENDIX for further details on the simulation program).

Analysis of NH+4-induced acidification in MD cells. First, average records were obtained from five experiments in which pHi was measured following luminal NH+4 addition in control conditions or in the presence of different inhibitors. Acceptable fits could be obtained with a variety of PaNH3 values ranging from 10 to 40 s-1. Interestingly, in this range of PaNH3, good fits could be obtained with PaNH3 = PblNH3. For example, the fits shown in Fig. 5, A-D, were obtained with the parameters given in Table 2, in which PaNH3 and PblNH3 were set equal to 20 s-1. All the characteristics of the recorded pHi values are correctly reproduced by the model, and none of the remaining parameters (PaiNH4, PeNH4, SH) could be changed by more than 25% without sensibly affecting the quality of the fits. The simulation program used with the best set of parameters (with PaNH3 = PblNH3 = 20 s-1), which is given in Table 2, displays unexpected features. First, the large NH+4-induced acidification rate observed in control conditions can be closely reproduced with an apical NH3 permeability 30 times larger than the apical NH+4 permeability for influx (this observation is valid for the whole range of acceptable PaNH3 values). This relatively large ratio of NH3 to NH+4 permeabilities contradicts previous estimations for MTAL where, with pHi recordings quite similar to those presented here, PaNH3 was assumed low or negligible with respect to PaiNH4 (12, 13, 27). In rat MTAL, the apical NH3 permeability was latter shown to exist in experiments where NH+4 pathways were blocked (8). Experimentally, this significant PaNH3 in MD cells expresses itself by a small initial alkalinization upon NH+4 addition, which was, in some occasions, clearly observed (see Figs. 2B and 3), and by a larger acidification upon washing out luminal NH+4. A second feature readily explained by the model is the experimental observation that pHi recovery after NH+4 removal is insensitive to bumetanide and Ba2+. The model reveals that it takes only 4.5 s at the beginning of the washout period to bring the intracellular NH+4 concentration from 38 mM at the end of the luminal NH+4 application to 1 mM which causes the rapid acidification observed in the first few seconds following NH+4 removal. The relatively slower pHi recovery following this initial acidification is the expression of a classic pHi regulation system mainly depending on the Na+/H+ exchanger. A third feature of the best fit obtained is that the initial NH+4 influx is not 40 times larger than the proton production rate as predicted earlier (27) but only approx 2.1 times larger (JNH4 = 13 mM/s and dpHi/dt × beta i = 6.3 mM/s). Even if JNH4 is not equal to the proton production rate, comparison of Table 1 and 2 shows that the JNH4 values calculated by the model are in proportion with the measured dpHi/dt values in the different experimental conditions.


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Fig. 5.   Fits of the average pHi time courses following luminal addition of 20 mM NH+4. Parameters in the transport model are given in Table 2 for the following four cases studied: control (A), in presence of 5 mM Ba2+ (B), in presence of 5 µM bumetanide (C), and in presence of both inhibitors simultaneously (D).

                              
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Table 2.   Parameters used for the different curves of Fig. 5

Absolute value of apical NH+4 influx. Beeuwkes and Rosen (3) have suggested a low or absent Na+-K+-ATPase activity in MD cells, and microenzymatic measurement of Na+-K+-ATPase activity revealed an enzyme activity per unit of cell volume of ~1/40th of that found in TAL (25). Low levels of Na+-K+-ATPase in the basal membrane of MD cells have also been demonstrated by using monoclonal antibodies against the enzyme (11). Thus those studies suggest that relative to TAL cells, MD cells cannot maintain a very significant transepithelial NaCl transport through, presumably, the activity of the apical Na+:K+:2Cl- cotransporter and a Ba2+-sensitive pathway and a basolateral Na+-K+-ATPase. It is therefore surprising to find in the present study an NH+4-induced proton production rate of comparable amplitude vs. that found in TAL. If the MD apical permeability coefficient for NH+4 was in proportion to the low activity expected for the basolateral Na+-K+-ATPase (everything being normalized to the cell volume), then the NH+4-induced proton production rate (also per unit of cell volume) would be expected to be at least one order of magnitude lower for MD cells vs. TAL cells.

Transport pathways for luminal NH+4. One of the pathways used by NH+4 to induce bumetanide-sensitive cell acidification is very likely the Na+:K+(NH+4):2Cl- cotransport. We found that 5 µM bumetanide inhibits 57% of the initial rate of cell acidification observed with luminal addition of NH+4. This finding agrees with recent work which indicated that the active NH+4 transport in TAL proceeds via the substitution of NH+4 for K+ in the apical membrane Na+:K+:2Cl- cotransporter (1, 8, 12-14, 27).

Our studies show that bumetanide did not completely prevent the intracellular acidification induced by luminal NH+4 and that the residual acidification (35-40% of control) observed with bumetanide was inhibited by luminal Ba2+. On the basis of previous observations that a component of NH+4-induced cell acidification was Ba2+ sensitive, it was suggested that NH+4 entry in the TAL occurs via apical membrane K+ channels (12, 13, 27). We have indeed directly observed a single class of K+ channels in the apical membrane of MD cells using the patch-clamp technique (9). However, we have also shown that complete inhibition of these K+ channels can be achieved by increasing intracellular [Ca2+] with 1 µM ionomycin. As this maneuver does not prevent the Ba2+-sensitive NH+4-induced cell acidification (Fig. 3), it is unlikely that NH+4 ions use the K+ channels that we have previously observed (9). We have to recognize that a different class of K+ channels that may have remained undetected in our patch-clamp experiments could be responsible for the Ba2+-sensitive NH+4 influx observed in the present study. Luminal application of 5 mM Ba2+, however, produces an instantaneous cell acidification (see Fig. 2C) that is unexpected from the blockade of K+ channels and the ensuing depolarization. It was recently argued that for rat MTAL, the Ba2+-sensitive portion of NH+4-induced acidification was not related to K+ channels but rather to a verapamil-sensitive K+/H+(NH+4) antiport (1). This is not likely to be the case in MD cells as, first, 0.1 mM verapamil had no effect on NH+4-induced acidification, and second, contrary to what was happening in TAL cells, Ba2+ systematically produced an instantaneous acidification of MD cells, which discards any direct effect from a putative K+/H+ exchanger in MD cells. The nature of this NH+4 pathway could not be identified in the present studies; however, the transporter/channel involved should mediate H+ efflux under control conditions, be Ba2+ sensitive, and transport NH+4 from the lumen to the cytosol upon luminal NH+4 addition.

In conclusion, MD cells behave just like MTAL cells when 20 mM NH+4 is presented in the luminal perfusate: a dominant cell acidification is observed with bumetanide-sensitive and Ba2+-sensitive components. Contrary to previous interpretations, these results can be quantitatively explained with an apical NH3 permeability 30 times larger than the NH+4 apical permeability, if we allow for a significant basolateral NH3 permeability and/or an apical NH+4 entry mechanism which has the capacity to accumulate NH+4 in the cell. The calculated NH+4 apical permeability is surprisingly high with respect to MTAL, given the fact that the basolateral pumping capacity was reported to be much lower.

    APPENDIX
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Abstract
Introduction
Materials & Methods
Results
Discussion
Appendix
References

Transport model for simulating NH3/NH+4 fluxes in epithelial cells. pHi, [NH+4], and [NH3] can be simulated on a personal computer as a function of time following luminal addition of 20 mM NH+4. Concentrations are given in units of mM, permeabilities in units of s-1, and fluxes appear in units of mM/s as defined in MATERIAL AND METHODS. As briefly explained in the DISCUSSION, the following five parameters are used in this simple model: apical and basolateral permeability coefficients for NH3 (PaNH3, PblNH3), permeability coefficients for NH+4 entry across apical membrane (PaiNH4) and for NH+4 exit across both apical and basolateral membranes (PeNH4), and the sensitivity (SH) of a pHi regulatory mechanism which transports protons out of the cell according to the simple relation JH = SH × (7.5 - pHi). For each time increment (0.1 s), unidirectional influx and efflux of NH+4 and NH3, and resulting intracellular concentrations are calculated. Then, at a given pHi, [NH+4] is allowed to equilibrate with [NH3] and [H], keeping the total [NH+4] + [NH3] constant. This equilibration process releases or captures a given amount of protons which are added to the efflux of proton mediated by the pHi-regulating mechanism. Finally, a new pHi is calculated based on the proton net flux and the known buffer capacity of the cell.

Each of the five parameters play a specific role, as can be easily seen in the simulation. SH, the sensitivity of the pHi regulatory mechanism, has, of course, a crucial role to play in the amplitude of the acidification produced by apical NH+4 addition. While other parameters can also influence this amplitude, SH is the only parameter affecting the slow alkalinization observed in the recovery period following removal of apical NH+4. PaNH3 specifically influences the size of the initial pHi changes upon apical NH+4 addition or removal. At a given SH and PaNH3, PaiNH4 affects the NH+4-induced acidification rate, and the ratio PaiNH4/PeNH4 together with PblNH3 determine the final pHi reached. This makes good sense as the pHi level reached in the presence of apical NH+4 depends on a large intracellular proton production rate. This proton production rate is maintained elevated if the dissociation reaction NH+4 = NH3 + H+ is set slightly off equilibrium by a concentrating NH+4 uptake mechanism (large PaiNH4/PeNH4) or by a low [NH3] set by a large basolateral NH3 permeability coefficient (PblNH3). The model clearly shows that the large acidification rate seen after apical NH+4 addition is not indicative of a larger apical permeability for NH+4 than for NH3. If one sets PaiNH4 PeNH4, and if the NH3 efflux through the basolateral membrane is minimized (PblNH3 = 0), then one cannot generate a significant NH+4-induced cellular acidification no matter how large one sets the apical NH+4 permeability. The conditions that generate such an acidification rate are a reduction of [NH3] through basolateral exit and an accumulative mechanism for apical NH+4 entry, which can be secondary to cotransported solutes or to membrane electrical potential.

    ACKNOWLEDGEMENTS

This work was supported by the Kidney Foundation of Canada awarded to J.-Y. Lapointe.

    FOOTNOTES

Address for reprint requests: J.-Y. Lapointe, Groupe de Recherche en Transport Membranaire, Université de Montréal, P.O. Box 6128 Succursalle (centre-ville), Montréal, Quebec, Canada H3C 3J7.

Received 11 October 1996; accepted in final form 16 July 1997.

    REFERENCES
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
Appendix
References

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