Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut 06520
Submitted 17 March 2003 ; accepted in final form 15 June 2003
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ABSTRACT |
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intracellular pH; carbon dioxide; out-of-equilibrium solutions; fura 2; ions; transport; kidney
The rate of H+ secretion by the proximal tubule, which is nearly
identical to JHCO3, is under the control of several
hormones. For instance, angiotensin II
(36,
71) and nitric oxide
(70) increase
JHCO3, whereas parathyroid hormone (PTH) has the opposite
effect (21,
41). Another potent regulator
of JHCO3 is the acid-base status of blood. For example,
respiratory acidosis {i.e., an increase in blood PCO2 that causes a
decrease in blood pH and small increase in blood
concentration
(
)} raises
JHCO3 (1,
11,
22). To determine whether it
is a change in PCO2, pH, or
that is responsible for the
increase in JHCO3 during respiratory acidosis, the
laboratory developed a technique for making out-of-equilibrium (OOE)
solutions. Using this approach it is possible to generate solutions having
physiological levels of CO2 concentration ([CO2]) and pH
but virtually no
(i.e., a
"pure CO2" solution), or solutions having physiological
levels of
and pH but virtually
no CO2 (i.e., a "pure
" solution).
OOE solutions were first used to study K-HCO3 cotransport in
squid giant axons (75). More
recently, our laboratory adapted this technique to mammalian cells and found
that removing from the basolateral
or "bath" solution (pure CO2) caused
JHCO3 to increase, whereas removing CO2 from
the basolateral solution (pure
)
had the opposite effect (76).
In other experiments, our laboratory used the OOE approach to vary basolateral
[CO2],
, and pH one at
a time, while holding the other two parameters constant. The most surprising
result was that JHCO3 was totally insensitive to wide
changes in basolateral pH, even though these changes in basolateral pH were
associated with rather wide changes in intracellular pH (pHi).
Nevertheless, JHCO3 increased markedly in response to
increases in basolateral [CO2]
(77). These results led to the
hypothesis that renal proximal tubule cells have a mechanism at or near the
basolateral membrane for sensing CO2 independently of pH. This
hypothesis is consistent with earlier work with equilibrated
solutions that showed that adding
to the
bath, but not to the lumen, causes steady-state pHi to rise in
proximal tubule cells (46) and
stimulates luminal acid extruders
(17,
18).
In the present study, we investigated one of the potential intracellular
signaling pathways of the basolateral CO2 sensor by monitoring
intracellular Ca2+ concentration
([Ca2+]i). Intracellular
Ca2+ is a common second messenger for numerous stimuli
(19). For example, a rise in
[Ca2+]i is a key step in the response of the
chemosensitive cells in the carotid body to hypoxia, metabolic acidosis,
respiratory acidosis, or isohydric hypercapnia
(13). Based on these findings,
we felt that a rise in [Ca2+]i was a good
candidate as a signaling pathway for the CO2 sensor. Some authors
working on proximal tubule cells have reported that increases in
[Ca2+]i raise JHCO3
(38), whereas others have
reported that increases in [Ca2+]i lower
JHCO3
(16). Here, using a
Ca2+-sensitive fluorescent dye, we found that we could
trigger a significant increase in [Ca2+]i by
introducing equilibrated
to the
basolateral (but not luminal) side of the tubule, or by introducing
basolateral pure CO2 (but not pure
). Also, we found that basolateral
CO2 does not increase [Ca2+]i by
lowering pHi and that the source of the Ca2+
is a thapsigargin (Tg)-insensitive intracellular store. Our results are thus
consistent with the hypothesis that an increase in
[Ca2+]i might be involved in the proximal
tubule cell's response to basolateral CO2.
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METHODS |
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All the experiments were carried out in "pathogen-free" female rabbits (New Zealand White, Elite, Covance, Denver, PA) weighing 1.42.0 kg. The methods for preparing the animals, harvesting the kidneys, and perfusing the tubules were similar to those originally described by Burg et al. (14) and subsequently modified in our laboratory (47, 76). The Yale Animal Care and Use Committee approved all the procedures. Briefly, an animal was euthanized by intravenous injection of pentobarbital sodium; an incision of the abdominal wall was performed to expose the left kidney, which was rapidly removed. The kidney was then cut into coronal slices and kept in cold (4°C) modified Hanks' solution (solution 1 in Table 1). The microdissection of the slice was carried out in the same solution under a dissecting microscope, using a pair of fine forceps to grasp a portion of a medullary ray and gently tear it from the rest of the slice, starting from the inner medulla and proceeding toward the cortex. Our initial landmark was the junction between the thin descending limb of Henle's loop and the S3 segment (i.e., distal portion of the proximal straight tubule). We isolated a portion of the S2 segment that consisted of the distal-most 600800 µm of the proximal convoluted tubules plus 200300 µm of the proximal-most part of the proximal straight tubule. After transferring the tubule to a chamber (adapted for rapid mixing of OOE solutions; see below), we perfused the distal-most 400500 µm of the proximal convoluted tubule. Tubules were perfused and bathed at 37°C.
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Solutions
The compositions of the solutions are given in
Table 1. The HEPES-buffered
solution (solution 2), the equilibrated
solution (solution 3), the pure CO2 OOE solution
(solution 4), and the pure
OOE solution (solution 5)
were adjusted to pH 7.40 at 37°C. The osmolalities, measured using a
vapor-pressure osmometer (model 5100C, Wescor, Logan UT), were adjusted to 300
± 3 mosM. The method for generating OOE solutions was the same as
originally described (75), as
adapted for kidney tubules
(76). Briefly, we generated
OOE solutions by exploiting the slow interconversion between CO2
and
to generate 20% pure
CO2 (i.e., virtually no
) and rapidly mixed solution
4/part A and solution 4/part B
(Table 1), each contained in a
140-ml plastic syringe (140 ml, Monoject, Sherwood Medical Industries,
Ballymoney, UK) driven by the same syringe pump (model 552222, Harvard
Apparatus, South Holliston, MA). The output of the syringe was connected to an
array of five-way valves (Eagle P/N E41PP-00000, Clippard
Instrument Laboratory, Cincinnati, OH) with Tygon tubing
(
in. ID, Norton Performance Plastics, Akron, OH), and then directed to
stainless steel tubing surrounded by a water jacket to warm the solution
sufficiently so that the temperature in the chamber was 37°C. Shortly
downstream from the water jacket, the output of the stainless steel tubing was
connected via Tygon tubing to a mixing "T," which in turn was
connected to another length of Tygon tubing that was filled with nylon mesh to
promote mixing. Finally, this Tygon tubing was connected to the chamber, which
consisted of a straight canal that was 14-mm long x by 2.5-mm wide to
promote a laminar flow. A comparable method was used to generate the pure
OOE solution. All solutions flowed
at 7 ml/min.
When using ionomycin for calibrating the Ca2+-sensitive dyes, or when using nigericin for calibrating the pH-sensitive dye (see below), we introduced these agents into the chamber via solution reservoirs, tubing, and an inlet port that were completely separate from those used for the physiological solutions. This precaution avoided contamination of the plumbing fixtures used for the physiological solutions. After each experiment, we washed the chamber extensively with 70% ethanol in water to remove traces of ionomycin or nigericin (5).
For solutions containing 0.5 mM ATP (Sigma, St. Louis, MO), we increased the total concentration of CaCl2 to 1.09 mM and the total concentration of MgSO4 to 1.57 mM to compensate for the binding of Ca2+ and Mg2+ to ATP. For solutions containing 0.5 mM EGTA, we increased the total concentration of MgSO4 to 1.26 mM. For solutions containing 5 mM EGTA, the total concentration of MgCl2 was increased to 1.82 mM to compensate for Mg2+ binding to EGTA. We used BAD computer software described by Brooks and Storey (12) to compute the amount of extra CaCl2, MgSO4, or MgCl2 that we needed to add to maintain the free Ca2+ at 1 mM and the free Mg2+ at 1.2 mM.
Chemicals. 4-Bromo A-23187, ionomycin, and Tg, were obtained from Calbiochem (Calbiochem-Novobiochem, La Jolla, CA). Rotenone was obtained from ICN (ICN Pharmaceuticals, Costa Mesa, CA). HEPES was obtained from USB (USB, Cleveland, OH). Nigericin, ATP, and the other chemicals in the physiological solutions were obtained from Sigma.
Fluorescence Measurements
Measurement of fluorescence of Ca2+-sensitive dyes. Measurements of [Ca2+]i were performed by loading the isolated perfused tubule with 5 µM of either fura 2-AM (Molecular Probes, Eugene, OR) or fura-PE3-AM (TefLabs, Austin TX) in our HEPES-buffered solution (solution 2 in Table 1) along with 0.5% (vol/vol) pluronic F-127 (Molecular Probes). We dye-loaded the tubules at room temperature for 2030 min for fura 2 and 4050 min for fura-PE3. We added the dye precursors as 2 mM stock solutions in DMSO and added the pluronic F-127 as a 20% wt/vol stock solution in DMSO. Before the fluorescence recordings, we washed the tubule by flowing a large volume of solution 2 through the chamber.
In tubules loaded with fura 2, dye leakage led to a gradual loss of fluorescence that often prevented us from performing longer experiments. Therefore, in lengthy experiments (>25 min), we used fura-PE3, which is more resistant to dye leakage, has the same absorbance spectrum as fura 2 (69), and has been used successfully in proximal tubule cells by others (53). However, we used fura 2 in most of our experiments because fura-PE3 required a longer period of dye loading, which reduced the number of experiments we could perform per rabbit and also increased our failure rate. Therefore, unless our experimental protocol required that we record [Ca2+]i for a lengthy period, we preferred fura 2 over fura-PE3.
The microscope was an Olympus IX70 inverted microscope, equipped with a x40 oil-immersion objective (1.35 numerical aperture, with a x1.5 magnification selector knob) and apparatus for epi-illumination. The light source was a 75-W xenon arc lamp. We generated light at two excitation wavelengths by using a filter wheel (Ludl Electronic Products, Hawthorne, NY) to alternate the placement of two filters, 340 ± 15 and 380 ± 15 nm (Omega Optical, Brattleboro, VT), in the excitation light path. Appropriate neutral-density filters (Omega Optical) mounted on a second wheel were used to avoid overillumination of the specimen, which could cause photobleaching, and to equalize as nearly as possible the emitted fluorescent light intensities obtained while excitation occurred at the two wavelengths. The excitation light was directed to the tubule via a 415-nm long-pass dichroic mirror (DM 415, Omega Optical) and the aforementioned objective. The emitted light was collected by the same objective and, via a band-pass filter (510 ± 40 nm, Omega Optical), was directed to an intensified CCD camera (model 350F, Video Scope International, Dulles, VA).
The protocol for alternately exciting the tubule with wavelengths of light,
and for subsequently acquiring the fluorescence images, was described
previously (76). Briefly, a
typical data-acquisition cycle consisted of a 370-ms period of
illumination with 340-nm light, followed immediately by an identical period
with 380-nm light. For each excitation wavelength, we averaged four successive
video frames using an image-processing board (DT3155, Data Translation,
Marlboro, MA) and thereby obtained the emitted light intensity for an
excitation of either 340 nm (I340) or 380 nm
(I380). This pair of excitations was repeated at intervals
ranging from 2.5 to 8 s; between excitations cycles, a shutter on the filter
wheel protected the tubule from the light. Software developed in our
laboratory using the Optimas (Media Cybernetics, Silver Spring, MD) platform
controlled data acquisition and analysis on an Intel-based computer running
Windows 98SE. We identified an area of interest that represented
30% of
the tubule length. The sum of the I340 values of the
pixels in the area of interest, corrected for the background (see below), was
divided by the sum of the corresponding background-subtracted
I380 values to yield the fluorescence excitation ratio
(I340)/(I380) or R340/380,
which strongly depends on [Ca2+]i but is
relatively insensitive to factors such as dye concentration. Because it was
our impression that sudden increases in the rate of dye loss were associated
with sudden increases in
(I340)/(I380), we discarded
experiments in which I340 and I380
declined rapidly.
Intracellular calibration of
Ca2+-sensitive dyes. The generally
accepted approach for converting R340/380 values into
[Ca2+]i values is that of Grynkiewicz et al.
(27), in which one determines
Rmin (the minimum R340/380 when
[Ca2+]i 0) and Rmax (the maximum
R340/380 when [Ca2+]i
) for
each cell, and computes [Ca2+]i on the
assumption that the dissociation of dye is the same inside the cell as it is
in vitro
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Unfortunately, as reported by several groups, the above calibration approach is difficult to apply to isolated proximal tubules because of problems with dye leakage during the prolonged calibration procedure. Thus most [Ca2+]i studies on proximal tubules, and the associated calibrations, have been done on collapsed tubules (9, 40, 74). Another group performed their physiological experiments in perfused tubules but obtained the values for Rmin, Rmax, and Sf/b by performing calibrations on collapsed tubules (4). We know only one study in which the authors calibrated a Ca2+-sensitive dye (i.e., fura-PE3) in a limited number of perfused proximal tubules (53).
Despite various attempts to minimize dye loss and cell damage, we found it impossible to perform physiological experiment and then routinely obtain Rmin, Rmax, and Sf/b values on the same perfused tubule at 37°C. For example, although probenecid (an inhibitor of organic anion transporters) reduces the loss of fura 2 from neurons (44), we did not find probenecid (3001,000 µM) useful in the proximal tubule. Similarly, neither lowering the ionomycin concentration to 1 µM, nor switching from ionomycin to 4-bromo A-23187 was helpful. Instead, we adopted the following procedure.
First, we obtained a mean Sf/b as well as mean, normalized values of Rmin and Rmax on a subset of 30 tubules perfused at 37°C after we had performed physiological experiments on these tubules. At the end of the experiment, we switched successively to bath solutions containing 1) 0 mM Ca2+ plus 5 mM EGTA and 5 µM of the Ca2+ ionophore ionomycin (solution 6 in Table 1)1, 2) 5 mM Ca2+ plus 5 µM ionomycin (solution 7)2, and 3) 5 mM Mn2+ (solution 8). This last maneuver allowed us to determine the autofluorescence of the tubule by quenching the fluorescence of the dye. We subtracted these quenched values of I340 and I380 from all respective I340 and I380 values in the experiment and used these background-subtracted values to compute R340/380 values for each data point. Finally, we identified a segment of data at the beginning of the experiment in which the R340/380 values were stable with the HEPES-buffered solution (solution 2) present in the lumen and bath, calculated the mean initial R340/380 value, and divided all R340/380 values in the experiment by this mean initial R340/380 value. The mean quotient during the calibration period with 0 mM Ca2+ was thus the normalized Rmin, and the mean quotient during the calibration period with 5 mM Ca2+ was the normalized Rmax. In the 30 tubules, Rmin was 0.63 ± 0.05, Rmax was 6.07 ± 0.65, and Sf/b was 3.33 ± 0.51.
Second, we used the above values of Rmin, Rmax, and Sf/b to compute [Ca2+]i values in each of our experiments, including the 30 described above. In each of these experiments, we normalized all R340/380 values to the mean initial R340/380 value obtained with the HEPES-buffered solution present in the lumen and bath (see above). We then used Eq. 1 to compute [Ca2+]i values at each time point, employing the aforementioned mean value of Sf/b, the mean normalized values of Rmin and Rmax, and a Kd for fura 2 of 224 nM (27) or a Kd for fura-PE3 of 290 nM (69).
Measurement of pHi. The ratiometric optical technique
used to measure pHi was similar to that described above for
[Ca2+]i. Briefly, isolated microperfused
tubules were loaded with the acetoxymethyl ester of the pH-sensitive dye
BCECF-AM (Molecular Probes) at 10 µM final concentration, dissolved in the
HEPES-buffered Ringer (solution 2 in
Table 1). The excitation
band-pass filters were centered at 440 ± 5 and 495 ± 5 nm (Omega
Optical). We also used a 510-nm long-pass dichroic mirror and a 530-nm
long-pass emission filter (Omega Optical). We identified areas of interest as
outlined above for the [Ca2+]i measurements,
subtracted the background (0.3% of the signal in BCECF-loaded tubules)
from the I440 and I490 values as
described previously (76), and
computed the time course of I490/I440.
We discarded experiments in which the rate constant for the decrease in the
I440 signal (k440) exceeded
0.05 min1
(6).
We computed pHi values from the
I490/I440 ratios using a variation of
the high-K+/nigericin technique
(66), in which one performs a
one-point calibration at pHi 7.00
(10). At the end of each
experiment, we drove pHi toward 7.00 by introducing a pH-7.00
high-K+/nigericin solution
(54) into the bath. We
normalized the I490/I440 ratios of the
entire experiment by dividing them by the
I490/I440 ratio obtained at
pHi 7.00 and then used the following equation
(10) to calculate
pHi
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From a separate series of 64 fluorescence measurements in a total of 10 tubules, we obtained values for pK and b by using a nigericin-containing solution to alter pHi, as described elsewhere (54). We used a nonlinear least-squares method to fit the parameters in the above equation, which forces the best-fit curve to pass through unity at pHi = 7.00, to the calibration data. The best-fit values were pK = 7.24 ± (SD)0.01 and b = 1.79 ± (SD)0.02.
Data Analysis and Statistics
Except for the curve fitting discussed above, all the values are means
± SE, with n being the number of observations. The statistical
significance of the data was assessed by two-tailed Student's t-tests
on paired or unpaired data as indicated, using the Analysis Toolpack of
Microsoft Excel. Mean steady-state [Ca2+]i
values were obtained by averaging [Ca2+]i
values over a period of 1 min.
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RESULTS |
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Effect on [Ca2+]i of Applying
Unilaterally
Our first approach in studying the effect of
solutions on [Ca2+]i was to measure
[Ca2+]i while exposing either the luminal or
the basolateral side of the tubule, but not both, to 5% CO2/22 mM
at a fixed extracellular pH of
7.40. A typical recording is shown in Fig.
1A. At the beginning of the experiment, we bilaterally
perfused the tubule with a solution buffered to pH 7.40 with HEPES
(solution 2). After we switched the luminal solution from one
buffered with HEPES to one buffered with 5%CO2/22 mM
(solution 3),
[Ca2+]i slowly drifted upward by a small
amount (segment ab). On the other hand, after we removed the
from
the lumen (bc) and then introduced the
-buffered solution to the bath, [Ca2+]i
increased to a new and substantially higher steady-state value (cd).
Switching back to the
bath solution caused [Ca2+]i to return close
to baseline (de). Figure
1B shows that we obtained the same result when we made
the luminal and basolateral solution changes in the opposite order.
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The histogram in Fig.
1C represents the mean paired changes in
[Ca2+]i
([Ca2+]i) elicited in eight tubules by
switching the solution in either the lumen (filled bar, corresponding to
segment ab in Fig.
1A and cd in
Fig. 1B) or the bath
(stippled bar, corresponding to segment cd in
Fig. 1A and
ab in Fig.
1B) from HEPES-buffer to 5% CO2/22 mM
.
[Ca2+]i was not statistically
significant when we applied
to the
lumen (P = 0.8) but was significant when we applied
to the
bath (P < 0.002).
Effect on pHi of Applying
Unilaterally
To test the hypothesis that the increase in
[Ca2+]i was caused by a change in
pHi, we repeated the above protocol while measuring pHi
in a total of 14 different tubules. Because we included lactate in our luminal
solutions to mimic the conditions in other parallel experiments in our
laboratory, we anticipated that the tubules would have a high initial
pHi. Previous work has shown that adding lactate to the lumen of
the salamander proximal tubule, or adding acetate to the lumen of the rabbit
S3 segment, raises pHi by 0.2 due to the coupled apical entry
of Na+ and monocarboxylate followed by the coupled exit of
H+ and lactate (or lactate/OH exchange) across the basolateral
membrane (45,
59). Indeed, in 9 of the 14
tubules, the initial pHi was relatively high (averaging 7.54
± 0.08). However, for unknown reasons, in the other five tubules, the
initial pHi in HEPES was lower (averaging 7.23 ±
0.07).3 Regardless of
whether the initial pHi in HEPES was high or low, introducing 5%
CO2/22 mM
to the lumen
caused a sustained decrease in
pHi.4 On
the other hand, the initial pHi in HEPES had a major impact on the
pHi response when we added
to the
bath. In the nine tubules with a high initial pHi, introducing 5%
CO2/22 mM
to the bath
caused a sustained acidification, whereas in the five other tubules with a
lower initial pHi, introducing 5% CO2/22 mM
induced an alkalinization. The
results are summarized in the Table
2. As noted in the DISCUSSION, the divergent response
to the addition of basolateral
is
consistent with observations made in other preparations.
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Thus in all of the experiments in which we monitored pHi,
introducing luminal
caused
an acidification; in all experiments in which we monitored
[Ca2+]i, introducing luminal
had no
effect. The relationship between
pHi and
[Ca2+]i was just the opposite in
two-thirds of the experiments in which we added
to the
bath. In 9 of 14 tubules in which we monitored pHi, introducing
basolateral
caused
an acidification, just as if we had added
to the
lumen. However, in all experiments in which we monitored
[Ca2+]i, introducing basolateral
caused
an increase in [Ca2+]i. Therefore, a change
in pHi cannot be the cause of the
[Ca2+]i increase elicited by basolateral
.
Effect on [Ca2+]i of Applying Pure
orPure CO2 Basolaterally
The above experiments ruled out a role for pHi in the increase
of [Ca2+]i elicited by bath
but
did not discriminate between bath CO2 and bath
. Next, we used OOE solutions to
investigate separately the effect of pure CO2 (solution 4)
and pure
(solution 5) on
[Ca2+]i. We chose to use 20% CO2
(nominally no
, pH 7.40) because
this basolateral PCO2 causes a substantially larger stimulation of
JHCO3 in the S2 proximal tubule than does 5%
CO2 (77). A
HEPES-buffered solution continuously perfused the lumen. As shown in
Fig. 2A, introducing
22 mM pure
to the bath caused, at
most, a trivial increase in [Ca2+]i
(segment ab), whereas introducing 20% pure CO2 always
caused a substantial and sustained increase in
[Ca2+]i
(Fig. 2A, segment
cde). Removing the pure CO2 solution caused
[Ca2+]i to decrease rapidly, but not all the
way to the baseline. In a total of 10 similar experiments
(Fig. 2B), pure
elicited a mean
[Ca2+]i of 7 ± 2 nM (n
= 10; P < 0.01) from a mean steady-state
[Ca2+]i of 76 ± 3 (n = 10).
This small [Ca2+]i increase could be the
result of a small CO2 contamination in our pure
solutions. On the other hand,
measurements with a CO2 electrode did not detect CO2 in
the pure
solutions exiting the
mixing T of our OOE apparatus. In the same tubules, pure CO2
elicited a much larger
[Ca2+]i = 62
± 17 nM (n = 10; P < 0.005) from a mean steady
state [Ca2+]i of 78 ± 2 (n =
10). These results support the hypothesis that it is basolateral
CO2, not
, that is
responsible for increasing [Ca2+]i in the
proximal tubule.
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One should recall that in Fig.
2A, [Ca2+]i did not
fully return to its baseline value after removal of bath pure CO2
In a series of 9 tubules distinct from the 10 discussed above, we exposed the
basolateral side of tubules to twin pulses of pure CO2, with a
delay of 5 min between pulses. The mean steady-state
[Ca2+]i before the first pulse was 57
± 1 nM. Because [Ca2+]i often did not
return to the initial baseline, the mean steady-state
[Ca2+]i before the second pulse was
significantly higher, 107 ± 19 nM (P < 0.03). The
[Ca2+]i elicited by the first pure
CO2 pulse was 85 ± 27 nM, whereas the
[Ca2+]i elicited by the second pure
CO2 pulse (starting from a higher baseline) was only 48 ±
14, a difference that is on the verge of statistical significance (P
= 0.052).
In six other experiments, we measured pHi while switching the
basolateral solution from HEPES (solution 2) to pure CO2
(solution 4). The mean pHi in bilateral HEPES was 7.23
± 0.02, whereas the mean pHi during bath exposure to pure
CO2 was 6.87 ± 0.08, a mean difference of 0.36 ±
0.09. Thus, even though the pHi decrease elicited by basolateral
pure CO2 was substantially less than that elicited by luminal
(0.36
vs. the values of 0.55 and 0.53 shown in
Table 2), basolateral pure
CO2 triggered an increase in
[Ca2+]i, whereas luminal
did
not. This result thus provides additional support for the hypothesis that it
is CO2 itself, and not the change in pHi, that is
responsible for the [Ca2+]i increase in our
experiments.
A technical question that arises is whether the large decrease in
pHi elicited by luminal
may
have affected the ability of fura 2 to report
[Ca2+]i. Although in their original paper
Grynkiewicz et al. (27)
reported that fura 2 is poorly pH sensitive, others have reported that
lowering the pH causes the Kd of fura 2 to increase
(34,
39). In our experiments, we
did not attempt to correct for this pH sensitivity of the
Kd because we did not simultaneously measure
pHi and Ca2+. Thus we probably underestimated
the rise in [Ca2+]i induced by the pure
CO2 solution in tubules with a relatively high initial
pHi.
Mechanism of the [Ca2+]i Increase Induced by Basolateral CO2
Effect of bilateral Ca2+-free solutions
on the CO2-induced
[Ca2+]i increase. We next
investigated the source of Ca2+ responsible for the
CO2-induced increase in [Ca2+]i.
Our first approach was to expose the tubule briefly to basolateral 20% pure
CO2, as in the second half of
Fig. 2A, first in the
presence and then in the absence of Ca2+.
Figure 3A shows such
an experiment. Initially, the lumen and bath contained a HEPES-buffered
solution (solution 2). A control pulse of 20% pure CO2
elicited a [Ca2+]i increase (segment
ab) that averaged 16 ± 2 nM (n = 8) and was partially
reversed in this experiment by removing the CO2 (bc). We
then switched the luminal solution to a variant of solution 2 in
which we omitted the Ca2+ and added 0.5 mM EGTA to
chelate trace amounts of Ca2+. This removal (point
c) reversed the slow upward drift in
[Ca2+]i and caused
[Ca2+]i to begin to decrease slowly. When we
then similarly removed Ca2+ from the bath (point
d), [Ca2+]i fell more rapidly
(de). Because exposing tubules to Ca2+-free
solutions for long periods (10 min) interfered with tubule integrity, we
challenged the tubule with a second CO2 pulse even as
[Ca2+]i continued to decline. We found that
the second 20% pure CO2 pulse, in the continued bilateral absence
of Ca2+, caused a
[Ca2+]i increase (ef) that averaged
13 ± 2 nM (n = 8) and was indistinguishable from the first
(P = 0.12). On removal of the bath pure CO2 solution,
[Ca2+]i fell (fg) to a value that
was lower than the value prevailing before we applied the CO2.
Reintroducing Ca2+ to the lumen and bath restored
[Ca2+]i to its initial level (gh).
Figure 3B summarizes
the mean
[Ca2+]i values in the
presence and absence of extracellular Ca2+.
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Effect of bilateral nifedipine on the CO2-induced
[Ca2+]i increase. In
experiments similar to that shown in Fig.
3A, we examined the effect of adding 10 µM nifedipine,
which blocks dihydropyridine-sensitive (L- and T-type)
Ca2+ channels
(62), to both the lumen and
the bath (not shown). We found that control 20% pure CO2 pulses
elicited a mean [Ca2+]i of 10 ±
3 nM (n = 6), a value that was not significantly different from the
[Ca2+]i of 13 ± 1 nM elicited
by 20% pure CO2 in the presence of bilateral nifedipine (P
= 0.4). The results of the experiments in this and the previous paragraph
indicate that an influx of extracellular Ca2+ is not
directly responsible for the CO2-induced increase in
[Ca2+]i.
Effect of Tg on CO2-induced [Ca2+]i increase. If CO2 causes the release of Ca2+ from an intracellular store, then blocking the reuptake of Ca2+ into this store ought to deplete the store and reduce the size of the CO2-induced increase [Ca2+]i. Tg is a well-known inhibitor of SERCA, the Ca2+ pump responsible for the uptake of Ca2+ into the sarco- and endoplasmic reticulum (52, 65). Figure 4A shows an experiment in which we tested the effect of Tg on the CO2-induced increase in [Ca2+]i. As a control, we first exposed the basolateral side of the tubule to 20% pure CO2, observing a reversible increase in [Ca2+]i (abc). Adding 2 µM Tg to the lumen and bath caused a transient rise in [Ca2+]i (point c), probably due to the decrease in Ca2+ reuptake into the stores, as has been observed for other cell types (48, 61, 65). Subsequently exposing the tubule to 20% pure CO2 in the continued presence of Tg induced a rise in [Ca2+]i that was actually somewhat greater than in the absence of the drug. As summarized in Fig. 4C for a total of 10 experiments, Tg produced a small but statistically significant increase in the CO2-induced increase in [Ca2+]i.
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To verify that Tg was indeed blocking the sarco-endoplasmic Ca2+ pump, we performed a positive control experiment in which we used extracellular ATP to activate the P2Y purinergic receptor and thereby release Ca2+ from Tg-sensitive stores (9, 74). As shown in Fig. 4B, basolateral ATP (0.5 mM) caused a very large but transient rise in [Ca2+]i, and Tg virtually eliminated this effect. As summarized in Fig. 4C for a total of eight similar experiments, the inhibition by Tg was statistically significant.
One might argue that a desensitization of the P2Y receptor may have been
responsible for the absence of a [Ca2+]i
increase during the second ATP pulse in
Fig. 4B, an effect
that would have led us to overestimate the blockade by Tg. We therefore
performed a separate series of experiments (not shown) in which we exposed
tubules to two ATP pulses (5 min apart) in the absence of inhibitors. The
first exposure of the basolateral side of the tubule to 0.5 mM ATP caused a
mean
[Ca2+]i of 99 ± 20 nM,
whereas the second induced a mean
[Ca2+]i of 110 ± 22 nM
(n = 7); this difference is not statistically significant (P
= 0.6).
Finally, we also performed two experiments (not shown) similar to the one in Fig. 4B, but in which, in the presence of Tg, we first pulsed the tubule with 0.5 mM ATP and then with 20% pure CO2. Even though ATP had a minimal effect, CO2 still elicited an increase in [Ca2+]i. The results of these three series of Tg experiments thus indicate that CO2 does not cause the release of Ca2+ from Tg-sensitive Ca2+ stores.
Effect of caffeine. To explore the possibility that a ryanodine
receptor might be involved in the CO2-induced increase in
[Ca2+]i, we assessed the ability of caffeine,
a well-known agonist of this receptor
(32,
78), to raise
[Ca2+]i in proximal tubule cells. In a total
of four experiments similar to the one shown in
Fig. 5, we exposed the proximal
tubule to 10 mM caffeine for 2 min. The mean
[Ca2+]i value measured before application of
caffeine was 57 ± 1 nM; adding caffeine caused a mean
[Ca2+]i of 1 ± 2 nM, a value
not statistically different from the baseline value (P = 0.8). On the
other hand, applying ATP always caused a transient increase in
[Ca2+]i. In the same four experiments, from a
mean baseline [Ca2+]i of 58 ± 1 nM,
adding ATP caused a mean
[Ca2+]i of
128 ± 29 nM. We conclude from these experiments that S2 proximal
tubules have no demonstrable ryanodine receptor activity and that it is
unlikely that these receptors play a role in the CO2-induced
increase in [Ca2+]i.
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Effect of rotenone on CO2-induced [Ca2+]i. To explore the possibility that CO2 causes the release of Ca2+ from the mitochondria, we examined the effect of rotenone on the CO2-induced [Ca2+]i increase. Our protocol was the same as for Tg (see Fig. 4A). Because rotenone blocks electron transport, we would expect that rotenone would cause Ca2+ to leak out of the mitochondria. Indeed, applying 4 µM rotenone caused the baseline [Ca2+]i to increase from 100 ± 14 to 155 ± 24 nM (P < 0.02, n = 5). Nevertheless, as summarized in Fig. 6, pulsing with 20% pure CO2 produced, if anything, a larger [Ca2+]i increase in the presence of rotenone than in its absence, although the difference was not statistically significant (P = 0.08, n = 5). An unavoidable complication in these experiments is that rotenone undoubtedly disturbed cellular energy metabolism. If these changes in energy metabolism did not affect the mechanism by which CO2 releases Ca2+ from internal stores, we would conclude that the mitochondria are not the source of the Ca2+ released in response to CO2.
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DISCUSSION |
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Influence of Initial pHi in HEPES on the pHi
Response to Bath
As noted in RESULTS and summarized in
Table 2, 9 of the 14 tubules
that we tested had a relatively high initial pHi in HEPES and
underwent a sustained acidification when we introduced
to the
bath. The other five tubules had a relatively low initial pHi and
underwent an alkalinization when we introduced
to the
bath. This dependence on the initial pHi is consistent with three
previous observations made using other preparations.
First, in the rabbit S3 segment, perfusing the lumen with a
monocarboxylate-free solution results in a relatively low initial
pHi. Under these conditions, adding basolateral
causes
a transient pHi fall followed by a large and sustained rise
(46), reflecting a three- to
fourfold stimulation of apical Na/H exchangers and H+ pumps
(17,
18) that overcomes the
acidifying influence of the basolateral Na-HCO3 cotransporter.
However, when the tubule lumen is perfused with acetate, the initial
pHi is relatively high, and adding basolateral
causes
a large and sustained fall in pHi
(46).5
Finally, in both hippocampal neurons
(58) and hippocampal
astrocytes (7), the effect of
adding
on steady-state pHi critically depends on the initial
pHi. The induced alkalinization is greatest at the lowest initial
pHi values and gradually falls off (or even reverses in the case of
the astrocytes) at progressively higher initial pHi values. A
general explanation for all three cases is that relatively high pHi
values stimulate acid loading but inhibit acid extrusion.
Basolateral CO2 Directly Triggers an Increase in [Ca2+]i
Ten years ago, Nakhoul et al.
(46), working with the rabbit
S3 proximal tubule (which always had a relatively low initial pHi
under the conditions of their experiments), showed that adding
to the
lumen causes a sustained pHi decrease that is presumably due to
1) the rapid diffusion of CO2 into the cell, followed by
2) the formation of H+ and
and the sustained basolateral exit
of
. On the other hand, they found
that adding
to the
bath induces only a transient pHi decrease, followed by a sustained
increase that is presumably due to an increase in net acid extrusion from the
cell. Indeed, Chen and Boron
(17,
18) showed that adding
equilibrated
to the
lumen had no effect on rates of apical Na/H exchange and H+
pumping, whereas adding the same solution to the bath increased these rates by
two- to fourfold. Here, we report a parallel observation: adding
to the
lumen never elicits a significant rise in
[Ca2+]i, whereas adding
to the
bath always triggers an increase in [Ca2+]i.
Thus it appears that basolateral, but not luminal,
produces several unique effects: 1) an increase in steady-state
pHi when the initial pHi is low, 2) an increase
in apical H+ extrusion, and 3) an increase in
[Ca2+]i.
One key question is whether it is CO2 or
that causes the above three
effects. Other work from our laboratory shows that it is specifically
basolateral CO2, and not basolateral
, that increases
JHCO3 (76,
77). In the present study, we
have made an additional parallel observation: a 20% pure CO2
solution in the bath can elicit a substantial rise in
[Ca2+]i, whereas a 22 mM pure
solution in the bath cannot. If
CO2, and not
, is indeed
the trigger, why is it that CO2 added to the lumen does not diffuse
across the apical membrane and through the cytoplasm to exert a measurable
effect at the basolateral membrane? We presume that, under the conditions of
such an experiment, the CO2 concentration near the basolateral
membrane is too low to produce a measurable stimulation of some sort of a
CO2 sensor.
The observation that it is basolateral CO2 and not
that triggers the increase in
[Ca2+]i does not distinguish between the
possibilities that basolateral CO2 1) acts directly on the
tubule to raise [Ca2+]i or 2) acts
indirectly by lowering pHi, which in turn leads to a rise in
[Ca2+]i. A precedent for the latter
hypothesis is that cytosolic acidification causes
[Ca2+]i to rise in gastric parietal cells
(67), platelets
(67), and cultured collecting
duct cells (60). On the other
hand, cytosolic acidification causes [Ca2+]i
to fall in squid giant axons
(3), and cytosolic
alkalinization causes [Ca2+]i to rise in both
HT 29 cells (48) and rat
pancreatic acinar cells
(61).
Did pHi indirectly control
[Ca2+]i in our experiments? Although an
isolated increase in bath [] causes
pHi to increase
(76), we found that switching
to a pure
solution causes only a
trivial increase in [Ca2+]i
(Fig. 2). Thus, if
pHi controls [Ca2+]i, it would
have to be a pHi decrease that causes
[Ca2+]i to rise. Indeed, switching to a pure
CO2 solution in the bath caused pHi to fall by
0.35
and consistently caused [Ca2+]i to increase,
apparently supporting the
pHi-[Ca2+]i hypothesis. However,
we found that adding
to the
lumen always causes pHi to fall by >0.5
(Table 2) but has no effect on
[Ca2+]i
(Fig. 1), ruling out the
pH-i[Ca2+]i hypothesis. Finally,
as noted in the presentation of Table
2 in RESULTS, introducing equilibrated
into
the bath caused pHi to decrease by
0.3 in 9 of 14 tubules
(i.e., the high-pHi tubules) but caused
[Ca2+]i to rise in 8 consecutive tubules. The
chance of randomly choosing eight consecutive high-pHi tubules is
only
3%. We conclude that a change in pHi is not the
intermediary through which CO2 raises
[Ca2+]i. This conclusion represents a third
parallelism between the CO2-induced increase in
[Ca2+]i and CO2-induced changes in
acid-base transport: In the proximal tubule, the CO2-induced
increase in JHCO3 does not occur via a decrease in
pHi (77).
Ca2+ Originates From an As Yet Unidentified Intracellular Pool
Two pieces of evidence indicate that the immediate source of the Ca2+ for the CO2-induced increase in [Ca2+]i is an intracellular store. First, the CO2-induced increase in [Ca2+]i occurs even when Ca2+ is absent from the lumen and bath (Fig. 3). Second, the dihydropyridine derivative nifedipine fails to attenuate the CO2-induced increase in [Ca2+]i. We chose nifedipine because the proximal tubule has dihydropyridine-sensitive Ca2+ channels that mediate Ca2+ influx during volume regulation after a hypotonic shock (40), in response to PTH (63), or during hypoxia (49).
One of the classic types of Ca2+ stores in cells is the Tg-sensitive store, which often is triggered by inositol 1,4,5-trisphosphate (IP3). Indeed, the P2Y purinergic receptor on the basolateral membrane of the proximal tubule releases Ca2+ from a Tg-sensitive pool (9, 74). Although we confirmed that adding Tg blocks the rise in [Ca2+]i stimulated by extracellular ATP (Fig. 4B), we found the drug to be ineffective in reducing the magnitude of the [Ca2+]i increase elicited by basolateral pure CO2 (Fig. 4A). In fact, in the presence of Tg, a pure CO2 pulse elicits a greater [Ca2+]i increase than a matched pulse in the absence of the drug (Fig. 4C). It is possible that, with Tg preventing the loading of Tg-sensitive stores, Tg-insensitive stores may accumulate extra Ca2+ that they release in response to CO2, resulting in a larger-than-normal CO2-induced increase in [Ca2+]i.
Ca2+ pools released by the ryanodine receptor are usually also Tg sensitive. However, we ruled out the possibility that ryanodine receptors are involved in the CO2-induced release of Ca2+ by demonstrating that millimolar concentrations of caffeine, which lead to a Ca2+-independent activation of the ryanodine channel (32, 78), do not elicit a rise in [Ca2+]i in the proximal tubule.
One Tg-insensitive Ca2+ pool is the mitochondria (23, 26, 29). However, our rotenone data are not consistent with the hypothesis that CO2 causes the release of Ca2+ from mitochondria. Thus our data are consistent with the hypothesis that, via a CO2 sensor at or near the basolateral membrane, CO2 triggers the release of Ca2+ from a nonconventional intracellular store.
Other investigators have demonstrated that multiple nonmitochondrial Ca2+ stores, functionally and spatially distinct, may coexist in the same cell (25, 43, 50, 52) and have in particular demonstrated the presence of Tg-insensitive pools. For example, a variety of cell lines have a nonmitochondrial pool that can take up Ca2+ after maximal inhibition by Tg (51, 64). In goldfish somatotrophs, GnRH causes a release of Ca2+ from a Tg-insensitive store (30). Moreover, in sea urchin eggs, the second messenger nicotinic acid adenine dinucleotide causes the release of Ca2+ from a Tg-insensitive store that is distinct from that triggered by either IP3 or cADP-ribose (24, 35). The Ca2+ pumps responsible for accumulating Ca2+ in the Golgi apparatus are Tg insensitive. Certain agonists (e.g., arginine vasopressin, histamine) coupled to the generation of IP3 can partially release Ca2+ from this pool (43, 50). Thus several pools are candidates for the CO2-induced release of Ca2+.
Potential Roles of the CO2-Induced Increase in [Ca2+]i
Previous work has established conflicting precedents for the effects that increases in [Ca2+]i have on acid-base transport in the proximal tubule. Four lines of evidence suggest that an increase in [Ca2+]i is associated with an increase in acid-base transport and/or JHCO3. First, in experiments on in vivo microperfused proximal tubules, raising [Ca2+]i by the luminal addition of the Ca2+ ionophore A-23187 increases JHCO3 in a dose-dependent manner (38). Second, adding angiotensin II to the basolateral side of a proximal tubule leads to increases in both JHCO3 (37) and [Ca2+]i (31). Third, carbachol triggers an increase in [Ca2+]i (42, 56) and stimulates the Na-HCO3 cotransporter; conversely, the Ca2+ chelator BAPTA prevents the stimulation of the cotransporter (56). Fourth and finally, CO2 causes insertion of vesicles containing H+ pumps into the apical membrane of the proximal tubule (57). In the turtle bladder, the application of CO2 triggers a rise in Ca2+ (15), and this rise in [Ca2+]i is required for the apical insertion of vesicles (68). A similar process may be at work in the rabbit outer medullary collecting duct (28).
Three lines of evidence suggest that an increase in [Ca2+]i is associated with a decrease in acid-base transport and/or JHCO3 in the proximal tubule. First, increasing [Ca2+]i by adding ionomycin to the bath leads to a decrease in JHCO3 (16). Second, PTH, a potent inhibitor of JHCO3 (20, 21), also increases [Ca2+]i (63). And third, a rise in [Ca2+]i inhibits the apical Na/H exchanger (72, 73).
One explanation for the apparently divergent data discussed above is that the relevant changes in [Ca2+]i occur within microdomains, and local changes in [Ca2+]i are more important than global ones (25, 33, 55). Another explanation for these divergent effects is that they are the consequence of different frequencies of Ca2+ spikes or waves. In the context of these possibilities, it is difficult to predict the role that [Ca2+]i plays in the response of the proximal tubule to basolateral CO2. We propose that CO2 binds to a CO2 sensor at or near the basolateral membrane and, independently of a change in pHi, triggers the release of Ca2+ from a nonmitochondrial intracellular store that is insensitive to Tg. The released Ca2+ might 1) modulate cellular processes not directly related to JHCO3, 2) be part of a signal-transduction pathway that results in an increase in JHCO3, or 3) be part of a braking mechanism that helps prevent runaway JHCO3 during CO2 stimulation.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 In some experiments, we also switched the luminal solution to one
containing 0 mM Ca2+ plus 5 mM EGTA; this precaution
caused I340 and I380 to change more
rapidly but did not affect the steady values.
2 In some experiments, we reversed the order in which we applied the 5 and 0
mM Ca2+ solutions.
3 One reason for this relatively low pHi is that these tubules
could have relatively low activities of either the apical or basolateral
monocarboxylate transporters.
4 CO2 passively enters the cell across the apical membrane,
leading to the intracellular formation of H+ and
. Presumably, pHi fails
to recover from this acid load because the unstimulated apical Na/H exchanger
and H+ pump are unable to recover from the acidifying influence of
that exits across the basolateral
membrane via the elctrogenic Na-HCO3 cotransporter.
5 The cause of this acidification has not been investigated. One possible
explanation is that the basolateral Na-HCO3 cotransporter is more
active at a higher pHi and thus exerts a larger acidifying
influence. In addition, adding basolateral
might
inhibit monocarboxylate transport. The gain of acid-loading capacity and/or
the loss of acid-extruding capacity might overwhelm the alkalinizing effect of
stimulating the apical Na/H exchanger and H+ pump.
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REFERENCES |
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