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
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ABSTRACT |
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
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INTRODUCTION |
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.
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MATERIALS AND METHODS |
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 (
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.
where
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
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
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.
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RESULTS |
Buffering capacity. Intrinsic
buffering power was obtained as described above using 10 mM TMA. As
displayed in Fig. 1, one can see that
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
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.
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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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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+.
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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.
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DISCUSSION |
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
i as a
function of pHi in MD cells that
is obvious from the raw data (see
inset of Fig. 1). The average
i of MD cells at a
pHi of 7.5 is 70 mM/pH unit. In
mouse MTAL cells,
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
i values were
estimated for rat MTAL (85 mM/pH unit, unpublished data cited in Ref.
27) whereas in rabbit proximal tubules (16),
i was estimated to 42.8 mM/pH
unit (84.6 mM/pH unit in the presence of
CO2/
). In consequence, net proton production rates after adding luminal NH+4
(dpHi/dt ×
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
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 ×
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
2.1 times
larger (JNH4 = 13 mM/s and
dpHi/dt ×
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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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 |
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.
 |
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