An increase in intracellular calcium concentration that is induced by basolateral CO2 in rabbit renal proximal tubule

Patrice Bouyer, Yuehan Zhou, and Walter F. Boron

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


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Working with isolated perfused S2 proximal tubules, we asked whether the basolateral CO2 sensor acts, in part, by raising intracellular Ca2+ concentration ([Ca2+]i), monitored with the dye fura 2 (or fura-PE3). In paired experiments, adding 5% CO2/22 mM (constant pH 7.40) to the bath (basolateral) solution caused [Ca2+]i to increase from 57 ± 3 to 97 ± 9nM(n = 8, P < 0.002), whereas the same maneuver in the lumen had no effect. Intracellular pH (pHi), measured with the dye BCECF, fell by 0.54 ± 0.08 (n = 14) when we added to the lumen. In 14 tubules in which we added to the bath, pHi fell by 0.55 ± 0.11 in 9 with a high initial pHi, but rose by 0.28 ± 0.07 in the other 5 with a low initial pHi. Thus it cannot be a pHi change that triggers the [Ca2+]i increase. Introducing to the bath an out-of-equilibrium (OOE) solution containing 20% CO2/no caused [Ca2+]i to rise by 62 ± 17 nM (n = 10), whereas an OOE solution containing 0% CO2/22 mM caused only a trivial increase. Removing Ca2+ from the lumen and bath, or adding 10 µM nifedipine (L- and T-type Ca2+-channel blocker) or 2 µM thapsigargin [sarco-(endo) plasmic reticulum Ca2+-ATPase inhibitor] or 4 µM rotenone (mitochondrial inhibitor) to the lumen and bath, failed to reduce the CO2-induced increase in [Ca2+]i. Adding 10 mM caffeine (ryanodine-receptor agonist) had no effect on [Ca2+]i. Thus basolateral CO2, presumably via a basolateral sensor, triggers the release of Ca2+ from a nonconventional intracellular pool.

intracellular pH; carbon dioxide; out-of-equilibrium solutions; fura 2; ions; transport; kidney


A MAJOR ROLE OF THE KIDNEY is to maintain the pH of the extracellular fluid within normal limits. The proximal tubule actively participates in this activity by reabsorbing ~80% of the NaHCO3 filtered at the glomeruli and also by generating "new" to neutralize non-volatile acids generated by metabolism. Bicarbonate reabsorption (JHCO3) occurs as the apical Na/H exchanger and H+ pump secrete H+, and as this acid titrates luminal to CO2 and H2O under the influence of apical carbonic anhydrase (2, 20). The newly formed CO2 and H2O then diffuse into the cells, where soluble carbonic anhydrase regenerates H+ and in the cytosol. Finally, the aforementioned H+ extruders recycle H+ to the lumen, while the basolateral Na-HCO3 cotransporter moves to the blood (8). The generation of new is similar to the reabsorption of except that the H+ secreted into the lumen titrates a buffer (e.g., ) other than , and the intracellular CO2 and H2O derive from the blood rather than the luminal fluid.

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.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Biological Preparation

All the experiments were carried out in "pathogen-free" female rabbits (New Zealand White, Elite, Covance, Denver, PA) weighing 1.4–2.0 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 600–800 µm of the proximal convoluted tubules plus 200–300 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 400–500 µm of the proximal convoluted tubule. Tubules were perfused and bathed at 37°C.


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Table 1. Physiological solutions

 

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 55–2222, Harvard Apparatus, South Holliston, MA). The output of the syringe was connected to an array of five-way valves (Eagle P/N E4–1PP-00–000, 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 20–30 min for fura 2 and 40–50 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 -> {infty}) 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

(1)
where Sf/b is the I380 measured when [Ca2+]i -> 0 divided by I380 when [Ca2+]i -> {infty}. Rmin is typically determined by exposing the cell to a Ca2+-free solution containing EGTA and a Ca2+ ionophore. Similarly, Rmax is typically determined by exposing the cell to a solution containing a high concentration of Ca2+ plus the ionophore.

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 (300–1,000 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

(2)

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.


    RESULTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In 30 tubules in which we obtained individual Rmin, Rmax, and Sf/b values in each tubule and used these values to compute [Ca2+]i in each tubule, the mean, steady-state [Ca2+]i was 63 ± 10 nM. As discussed in METHODS, we also used the data from the above 30 tubules to compute mean values for Rmin, Rmax, and Sf/b and then used these mean calibration parameters to compute initial [Ca2+]i values in a total of 131 tubules, including the 30 noted above. The average steady-state [Ca2+]i for these 131 tubules was 61 ± 1 nM (n = 131), which is not significantly different from the mean value for the 30 tubules (P = 0.8, unpaired t-test).

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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Fig. 1. Effect on intracellular Ca2+ concentration ([Ca2+]i) of basolateral vs. luminal exposure to 5% CO2/22 mM . A and B: representative recordings of [Ca2+]i. An S2 proximal tubule loaded with the Ca2+-sensitive dye fura 2 was initially perfused in lumen and bath with a HEPES-buffered, -free solution (solution 2, Table 1). At the indicated times, the lumen and the bath solution were switched to a similar solution buffered with (solution 3). A and B differ only in the order of the solution changes. C: data summary. Filled bar, the mean change in [Ca2+]i ({Delta}[Ca2+i]) caused by the luminal switch from the HEPES buffer to ; stippled bar, comparable {Delta}[Ca2+]i for the corresponding basolateral solution change. The statistical analyses summarized in the figure are the results of paired 2-tailed Student's t-tests. The mean initial [Ca2+]i values with the HEPES-buffered solutions in the lumen and bath were 61 ± 4 nM just before the luminal switch to the -buffered solutions, and 57 ± 3 nM just before the corresponding basolateral switch; the difference between these values is not statistically significant (P = 0.5, paired 2-tailed Student's t-tests).

 

The histogram in Fig. 1C represents the mean paired changes in [Ca2+]i ({Delta}[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 . {Delta}[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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Table 2. Effect of 5% CO2/22 mM on intracellular pHi

 

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 {Delta}pHi and {Delta}[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 {Delta}[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 {Delta}[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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Fig. 2. Effect on [Ca2+]i of basolateral exposure to a 22 mM "pure " vs. a 20% "pure CO2" solution. A: representative recording of [Ca2+]i. The lumen and bath were initially perfused with a HEPES-buffered, -free solution (solution 2, Table 1). At the indicated times, the bath solution was switched to either a 22 mM pure solution (solution 5) or to a 20% pure CO2 solution (solution 4). B: data summary. Hatched bar, mean {Delta}[Ca2+]i caused by the basolateral switch from the HEPES buffer to pure ; open bar, comparable {Delta}[Ca2+]i for the corresponding switch to pure CO2 solution. The statistical analyses summarized in the figure are the results of paired 2-tailed Student's t-tests. The mean initial [Ca2+]i values with the HEPES-buffered solutions in the lumen and bath were 76 ± 3 nM just before the basolateral switch to the pure solution, and 78 ± 2 nM just before the corresponding switch to the pure CO2 solution; the difference between these values is not statistically significant (P = 0.6, paired 2-tailed Student's t-tests).

 

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 {Delta}[Ca2+]i elicited by the first pure CO2 pulse was 85 ± 27 nM, whereas the {Delta}[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 {Delta}[Ca2+]i values in the presence and absence of extracellular Ca2+.



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Fig. 3. Effect of Ca2+ removal (+EGTA) on the [Ca2+]i increase elicited by 20% pure CO2. A: representative recording of [Ca2+]i. The lumen and bath were initially perfused with a HEPES-buffered, -free solution (solution 2, Table 1). At the indicated times, the bath solution was twice switched to a 20% pure CO2 solution (solution 4), the second time after Ca2+ was removed from both the lumen and bath. B: data summary. Open bar, mean {Delta}[Ca2+]i caused by the basolateral switch from the HEPES buffer to the pure CO2 solution in the presence of Ca2+; checkered bar, comparable {Delta}[Ca2+]i in the absence of Ca2+. The statistical analyses summarized in the figure are the results of paired 2-tailed Student's t-tests. The mean initial [Ca2+]i values with the HEPES-buffered solutions in the lumen and bath were 68 ± 3 nM just before the basolateral switch to the pure CO2 solution in the presence of Ca2+, and 62 ± 3 nM just before the corresponding switch in the absence of Ca2+; the difference between these values is statistically significant (P < 0.03 paired 2-tailed Student's t-tests).

 

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 {Delta}[Ca2+]i of 10 ± 3 nM (n = 6), a value that was not significantly different from the {Delta}[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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Fig. 4. Effect of thapsigargin (Tg) on the [Ca2+]i increase elicited by either 20% pure CO2 or ATP. A: representative recording of [Ca2+]i during a bath pure CO2 pulse. The lumen and bath were initially perfused with a HEPES-buffered, -free solution (solution 2, Table 1). At the indicated times, the bath solution was twice switched to a 20% pure CO2 solution (solution 4), the second time in the continued presence of 2 µM Tg. B: representative recording of [Ca2+]i during a bath ATP pulse. The protocol is the same as in A, except that we twice pulsed with ATP (solution 2 containing 0.5 mM ATP). C: data summary. Open bar, mean {Delta}[Ca2+]i caused by the basolateral switch from the HEPES buffer to the pure CO2 solution in the absence of Tg; checkered bar, comparable {Delta}[Ca2+]i in the presence of the drug; wavy and horizontally striped bars, comparable data for the ATP pulses. The statistical analyses summarized in the figure are the results of paired 2-tailed Student's t-tests. For the above 4 data sets, the mean initial [Ca2+]i values with the HEPES-buffered solutions in the lumen and bath, just before the basolateral switch to the pure CO2 solution and ATP, were 1) 75 ± 4 nM and 2) 76 ± 4 nM, and the mean values in presence of Tg just before the switch to the pure CO2 solution and ATP were 3) 68 ± 13 nM and and 4) 66 ± 20 nM. Neither the difference between 1 and 3 (P = 0.5) nor the difference between 2 and 4 (P = 0.6) is statistically significant.

 

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 {Delta}[Ca2+]i of 99 ± 20 nM, whereas the second induced a mean {Delta}[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 {Delta}[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 {Delta}[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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Fig. 5. Representative recording of [Ca2+]i during sequential bath exposures to 10 mM caffeine and 0.5 mM ATP. The tubule lumen was continuously perfused with a HEPES-buffered, -free solution (solution 2, Table 1). At the indicated times, the bath solution was temporarily replaced twice, first with solution 2 supplemented with 10 mM caffeine and the second time with solution 2 supplemented with 0.5 mM ATP. In a total of 4 such experiments, caffeine elicited a mean {Delta}[Ca2+]i (compared with paired baseline [Ca2+]i value just before the application of caffeine) of 1 ± 2 nM (P = 0.8, paired t-test), whereas ATP elicited a mean {Delta}[Ca2+]i of 128 ± 29 nM (P < 0.02, paired t-test).

 

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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Fig. 6. Effect of rotenone on the [Ca2+]i increase elicited by 20% pure CO2. Open bar, mean {Delta}[Ca2+]i caused by the basolateral switch from the HEPES buffer to the pure CO2 solution in the absence of rotenone; filled bar, comparable {Delta}[Ca2+]i in the presence of 4 µM rotenone. See the text for details. The statistical analysis summarized in the figure is the result of a paired 2-tailed Student's t-test.

 


    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
By increasing the rate at which they transport into the blood, the kidneys play an important role in the response to respiratory acidosis. Although work from our laboratory indicates that the trigger for increased transport is basolateral CO2 per se, rather than the accompanying acidosis, the underlying intracellular signals have not yet been resolved. We performed the present experiments to evaluate whether a rise in [Ca2+]i might be an element in one of the signaling pathways by which basolateral CO2 acts on renal proximal tubule cells.

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.


    DISCLOSURES
 
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Program Project Grant PO1-DK-17433. P. Bouyer was supported by Bourse Lavoisier du Ministère des Affaires Etrangère Française and the National Kidney Foundation.


    ACKNOWLEDGMENTS
 
The authors thank Duncan Wong for computer programming and for providing information-technology assistance and Dr. Barbara Ehrlich for helpful discussions.


    FOOTNOTES
 

Address for reprint requests and other correspondence: P. Bouyer, Dept. of Cellular and Molecular Physiology, School of Medicine, 333 Cedar St., PO Box 208026, New Haven CT, 06520-8026 (E-mail: Patrice.Bouyer{at}Yale.edu).

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. Back

2 In some experiments, we reversed the order in which we applied the 5 and 0 mM Ca2+ solutions. Back

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. Back

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. Back

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. Back


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