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Squid giant axons recover from acid loads by activating a Na+-driven Cl–HCO3 exchanger. We internally dialyzed axons to an intracellular pH (pHi) of 6.7, halted dialysis and monitored the pHi recovery (increase) in the presence of ATP or other nucleotides, using cyanide to block oxidative phosphorylation. We computed the equivalent acid-extrusion rate (JH) from the rate of pHi increase and intracellular buffering power. In experimental series 1, we used dialysis to vary [ATP]i, finding that Michaelis-Menten kinetics describes JH vs. [ATP]i, with an apparent Vmax of 15.6 pmole cm−2 s−1 and Km of 124 µM. In series 2, we examined ATPγS, AMP-PNP, AMP-PCP, AMP-CPP, GMP-PNP, ADP, ADPβS and GDPβS to determine if any, by themselves, could support transport. Only ATPγS (8 mM) supported acid extrusion; ATPγS also supported the -dependent 36Cl efflux expected of a Na+-driven Cl–HCO3 exchanger. Finally, in series 3, we asked whether any nucleotide could alter JH in the presence of a background [ATP]i of ~230 µM (control JH = 11.7 pmol cm−2 s−1). We found JH was decreased modestly by 8 mM AMP-PNP (JH = 8.0 pmol cm−2 s−1) but increased modestly by 1 mM ADPβS (JH = 16.0 pmol cm−2 s−1). We suggest that ATPγS leads to stable phosphorylation of the transporter or an essential activator.
Sodium-driven Cl–HCO3 exchange is the predominant acid-extrusion mechanism in axons from the squid Loligo pealei. The transporter, which is blocked by stilbene derivatives such as 4-acetamido-4′-isothiocyanatostilbene-2,2′-disulfonate (SITS) and 4,4′-diisothiocyanatostilbene-2,2′-disulfonate (DIDS), exchanges intracellular Cl− for extracellular Na+ and an -related species (Russell and Boron 1976; Thomas 1976, 1977; Boron and De Weer 1976a; Boron and Russell 1983). In the squid axon, this transporter has an absolute requirement for ATP (Boron and De Weer 1976a). In principle, the transporter could require ATP either as a fuel or as a cofactor. The fuel hypothesis, which predicts that the ATP is stoichiometrically hydrolyzed, seems unlikely inasmuch as the Na+ gradient has sufficient energy to support transport (Boron 1989). The final evidence against the fuel hypothesis is that internally dialyzing an axon with adenosine 5′-(γ-thio) triphosphate (ATPγS), rather than ATP, at low intracellular pH (pHi) supports transport, even after the subsequent washout of ATPγS (Russell and Boron 1976). However, after preactivation of the transporter with ATPγS at low pHi, raising pHi causes gradual inactivation.
Although these data rule out the fuel hypothesis, they do not address the issue of whether ATP is required to bind to the transporter (or an essential activator) or to phosphorylate it. In principle, this question could be approached by examining whether ATP analogues such as adenosine 5′-(β,γ-methylene)triphosphate (AMP-PCP, in which a methylene group links the β and γ phosphates) and adenosine 5′-(β,γ-imido)triphosphate (AMP-PNP, with an imido linkage)—which cannot serve as substrates for protein kinases—can support transport.
In the present study, we internally dialyzed squid axons to a low pHi (6.7), halted dialysis and exposed the axons to , and then computed the equivalent acid-extrusion rate (JH) from the rate of pHi recovery from this acid load. In one series of experiments, in which we used dialysis to vary [ATP]i, we found that simple Michaelis-Menten kinetics describes the dependence of JH on [ATP]i, with an apparent Jmax of 15.6 pmol cm−2 s−1 and an apparent Km of 124 µM. In a second series of experiments, we examined axons dialyzed with a variety of nucleotides to determine if any could support transport in the absence of ATP. We found that only ATPγS supported a statistically significant level of transport. Moreover, as expected for a Na+-driven Cl-HCO3 exchanger, ATPγS supported -stimulated 36Cl efflux. Finally, in a third series of experiments, we asked whether the nucleotides could alter JH in the presence of a background [ATP]i of ~230 µM (control JH = 11.7 pmol cm−2 s−1). We found that 8 mM AMP-PNP modestly decreased transport (JH = 8.0 pmol cm−2 s−1), whereas 1 mM ADPβS modestly increased it (JH = 16.0 pmol cm−2 s−1). Thus, the apparent affinity of Na+-driven Cl–HCO3 exchange for ATP is similar to that of other ATP-dependent processes, and no nucleotide other than ATPγS could support transport in the absence of ATP.
The experiments were conducted at the Marine Biological Laboratory (Woods Hole, MA). Technical details have been described previously (Boron and Russell 1983; Boron 1985; Boron and Knakal 1989). Briefly, a 3–4 cm length of giant axon (400–700 µm in diameter) was dissected from the squid L. pealei and horizontally cannulated in a chamber designed for internal dialysis (Brinley and Mullins 1967). Cellulose acetate dialysis tubing (outer diameter 140 µm) was threaded down the length of the axon and perfused with a dialysis fluid (DF) at ~2.1 µl/min. Also inserted into the axon through opposite cannulas were a pH-sensitive microelectrode (Hinke 1967) and an internal reference microelectrode (filled with 3 M KCl). Details on microelectrode construction, use of high-impedance electrometers and other devices to handle electrode signals, computer acquisition of data and computer control of the experiments have been described elsewhere (Boron and Russell 1983; Boron 1985).
The standard -free artificial seawater (ASW) had the following composition (in mM): 425.2 Na+, 12 K+, 3 Ca+, 52.5 Mg2+, 521 Cl−, 5 anionic form of [2-hydroxyethyl]-1-piperazine-propane sulfonic acid (EPPS) and 5 neutral form of EPPS (pK ~ 8.0), 0.1 EDTA=, 1 CN−. The –containing ASW was made by replacing 12 mM KCl with 12 mM KHCO3 and equilibrating with 0.5% CO2, 99.5% O2. Moreover, in this –containing ASW, the EPPS concentration was raised to 30 mM, with corresponding reductions in [Mg2+] and [Cl−] to keep osmolarity constant. All solutions were titrated to pH 8.0 using NaOH or HCl. Osmolarity was adjusted to 965–970 mOsm/kg using mannitol or H2O.
The standard DF for acid loading had the following composition (in mM): 0 Na+, 413.3 K+, 400 Cl−, 14 glutamate, 13.3 anionic form of (2–9N-morpholino)-ethanesulfonic acid (MES), 13.3 neutral form of MES and 0.5 phenol red. The fluid was adjusted to pH 6.7 with NMDG base or HCl. Osmolarity was adjusted to 965–970 mOsm/kg using glycine or H2O.
We varied the [total Mg2+] in the DFs according to the formula [nucleotide] + 3 mM = [total Mg2+]. Because the nucleotides bind Mg2+ in a ~1:1 ratio, this provision should maintain an approximately constant free [Mg2+]i. To achieve the desired final [total Mg2+] in the DF (i.e., 0, 3, 4, 7, 8 and 11 mM), we mixed Mg2+-free DF with DF containing 25 mM Mg2+.
An ATP-buffered DF containing 4 mM ATP and 3 mM PEPPA (Altamirano et al. 1995) was made by adding aliquots from stock solution containing 400 mM ATP as well as a stock solution containing 300 mM phosphoenolpyruvate (PEP) plus 300 mM phosphoarginine (PA) to an ATP-free stock DF. Similarly, DFs with a range of [ATP] values plus a constant 3 mM PEPPA were made by mixing a DF containing 4 mM ATP/3 mM PEPPA with an ATP-free DF containing 3 mM PEPPA. Finally, we generated ATP-free DFs with a range of [PEPPA] values by mixing an ATP-free DF containing 3 mM PEPPA with an ATP-free/PEPPA-free DF. All of the DFs used for the [ATP]i dose–response curve were titrated to a pH of 6.7. We obtained all nucleotides from Sigma Aldrich (St. Louis, MO). ATPγS, AMP-PNP, adenosine 5′-(α,β-methylene)triphosphate (AMP-CPP) and AMP-PCP were present at a final concentration of 8 mM in the DFs. ADP, adenosine 5′-(β-thio)diphosphate (ADPβS), guanosine 5′-(β,γ-imido)triphosphate (GMP-PNP) and guanosine 5′-(β-thio)diphosphate (GDPβS) were present at a final concentration of 1 mM.
We assayed ATP concentrations as described previously (Altamirano et al. 1988). Briefly, previously dialyzed axons were cut at both cannulated ends and placed on small pieces of Parafilm® (Alcan, Neenah, WI) using sharp forceps. The axoplasm was then squeezed from the axon using a device similar to a miniature paint roller. After rapidly weighing the Parafilm with the axoplasm, we transferred the Parafilm containing the axoplasm to a glass tube containing a stop buffer (0.3 N perchloric acid, subsequently neutralized with 0.25 M KOH, 0.15 M 3-[N-morpholino]propanesulfonic acid [MOPS] and 0.15 KCl) and stored the material until the time of assay. ATP was measured using the firefly luciferin-luciferase assay kit (LKB Wallach, Turku, Finland). The principle of the assay is based on the firefly luciferase-catalyzed oxidation of d-luciferin in the presence of an ATP-magnesium salt and O2. The ATP can be quantified by the amount of light produced.
As described previously (Boron and Knakal 1985), pHi data were acquired by computer, and rates of pHi recovery from acid loads (dpHi/dt) were determined from a linear curve fit to the data. Net JH is defined as the net efflux of H+ (or other acid) plus the net influx of (or other base) and computed as the product of dpHi/dt, total intracellular buffering power—assumed to be the sum of the closed-system buffering power computed for the DF and the open-system buffering power for (Roos and Boron 1981)—and volume-to-surface ratio.
Axons were prepared as described above and dialyzed with the appropriate DF for 60–90 min prior to changing to an identical DF containing 36Cl (specific activity 0.25 µCi/µmol of Cl−). The DF was the same as above but with 150 mM Cl− rather than 400 mM Cl− (glutamate replacing Cl−). The ASW superfusing the axon was then collected and each timed sample counted using a liquid scintillation counter to an error of <5% (Altamirano et al. 1995).
Values are reported as means ± standard errors. Groups of data were compared using one-way analysis of variance (ANOVA) followed by Dunnett’s post hoc analysis, using KaleidaGraph® (Synergy Software, Reading, PA).
Earlier experiments on intact (i.e., undialyzed) squid giant axons, acid-loaded by exposure to 5% CO2, had shown that addition of cyanide to ASW blocks the pHi recovery that we now know is mediated by the Na+-driven Cl–HCO3 exchanger (Boron and De Weer 1976a). Figure 1 shows an experiment on an axon that we internally dialyzed for ~60 min (segment ab) with a pH-6.7, Na+-free DF that contained 8 mM ATP—a concentration that we used in an earlier study of ATPγS (Boron et al. 1988)—plus 3 mM PEPPA. The axon was superfused with the standard -free ASW containing 1 mM CN− to block oxidative phosphorylation. When we halted dialysis (point b), intracellular pH stabilized and slowly increased as control of pHi returned to the axon (bc). Exposing the axon to 12 mM CO2 ASW resulted in a small, brief acidification (cd) caused by the influx of CO2. The pHi then recovered (i.e., increased) due to stimulation of Na+-driven Cl–HCO3 exchange (de). Addition of 0.5 mM SITS prevented any further pHi increase (ef), as is expected of this transporter.
To determine the dose dependence of the ATP requirement of the Na+-driven Cl–HCO3 exchanger, we performed experiments similar to that shown in Fig. 1, except that we dialyzed the axons with DFs designed to achieve a broad range of [ATP]i in the living axon. Because dialysis, by itself, is insufficient to reduce axoplasmic [ATP] to near-zero values—due to endogenous ATP production—we included 1 mM CN− in the ASW. When we varied the added [ATP] of the DFs from 0 to 4 mM, we added 3 mM of the phosphagens (i.e., ATP buffers) PEP and PA in order to “clamp” axoplasmic [ATP] following cessation of dialysis. Pyruvate kinase converts PEP to pyruvate, simultaneously converting ADP to ATP. Similarly, arginine kinase converts PA to arginine, also simultaneously converting ADP to ATP. Finally, at the end of each pHi experiment, we expressed the axoplasm and measured its [ATP]. Table 1 summarizes the directly measured ATP concentrations. Figure 2 summarizes the [ATP]i dependence of JH. Note that the [ATP]i values in this plot are the measured values from Table 1, not the nominal [ATP]DF. A nonlinear least-squares fitting procedure yielded an apparent Km of 124 ± 19 µM and an apparent Vmax of 15.6 ± 6.3 pmole cm−2 s−1.
To address the question of whether ATP is required for phosphorylation vs. binding, we surveyed eight nucleotide analogues for the ability to support transport at very low [ATP]i values. The protocol was identical to that for the axon represented in Fig. 1 except that the DF (1) was free of ATP and PEPPA and (2) contained either no nucleotide or one of the eight test nucleotide analogues. In the absence of any added nucleotide (i.e., actual [ATP]i = 11 µM), JH (i.e., background flux) was 1.0 ± 0.5 (Fig. 3, leftmost bar and top data row in Table 1). Confirming the observation from a previous study, ATPγS supported a significantly greater flux, which amounted to 16.0 ± 1.3 pmole cm−2 s−1. Because ATPγ may contain some ADP, we also examined the effect of ATPγS in the presence of diadenosine pentaphosphate (AP5A), a potent inhibitor of adenylate kinase (Lienhard and Secemski 1973), which catalyzes the reaction 2 ADP ATP + AMP. However, AP5A did not reduce the ability of ATPγS to support transport. None of the other nucleotide trisphosphate analogues increased JH significantly above background, as determined by ANOVA. Similarly, none of the nucleotide diphosphates supported Na-driven Cl–HCO3 exchange.
To address the possibility that the ATPγS effect on acid extrusion was the result of another acid-extrusion process, we measured 36Cl efflux from axons dialyzed at pH 6.7 with solutions that contained either no nucleotides, 4 mM ATP or 8 mM ATPγS. The results (Table 2) show that both the ATP- and ATPγS-treated axons exhibited a higher rate of unidirectional 36Cl efflux when exposed to 12 mM , whereas in the absence of nucleotide, exposure to 12 mM had no effect on 36Cl efflux. Previous work has shown that stilbenes block the entire -stimulated 36Cl efflux (Russell and Boron 1976; Boron and Russell 1983).
In a third series of experiments, we asked whether the same nucleotides summarized in Fig. 3 could alter JH in the presence of an intermediate [ATP]i that would allow us to detect either a stimulation or an inhibition of transport. In 12 experiments, dialysis with a pH-6.7 fluid containing 3 PEPPA but no added nucleotides produced a JH of 11.7 ± 0.8 (Fig. 4, leftmost bar and sixth data row in Table 1). The measured [ATP]i in this group of axons was 231 µM (Table 1)—about twice the apparent Km. We added nucleotides to this DF at either 1 or 8 mM. Dialysis with 8 mM—but not with 1 mM—AMP-PNP modestly reduced JH to 8.0 ± 0.6 pmol cm−2 s−1. On the other hand, 1 mM ADPβS modestly increased JH to 16.0 ± 0.5 pmol cm−2 s−1.
One of the first-described characteristics of the Na-driven/ Cl–HCO3 exchanger was its ATP dependence in the squid giant axon. Here, we have determined that the apparent Km for ATP is ~124 µM, more than an order of magnitude lower than the axon’s physiological [ATP]i of ~4 mM (Brinley and Mullins, 1967). Of course, we obtained this Km value under a very specific set of conditions, and it might very well depend on such parameters as pH and ion gradients. Nevertheless, 124 µM is within the range of values for other ATP-dependent processes in the squid axon and elsewhere. For example, in the squid axon, the apparent Km is 86 µM for the Na/K/Cl cotransporter (Altamirano et al. 1988), 250 µM for the Na–Ca exchanger (Di Polo 1977) and 200 µM for the Na–K pump (Beauge and Di Polo 1981).
A Na-dependent Cl–HCO3 exchanger would have a stoichiometry of two H+ equivalents extruded for each Cl− extruded. Given -dependent 36Cl efflux of ~4.6 ± 0.7 pmole cm−2 s−1 in the presence of 8 mM ATPγS (Table 2), we might have expected an equivalent H+ flux of ~9.2 pmole cm−2 s−1 if [Cl−]i were only 150 mM (as in the 36Cl experiments), but ~11.8 pmol cm−2 s−1 when we recall that [Cl−]i was in fact 400 mM in the pHi experiments. In fact, for the data in Fig. 2, in which ATP supported transport, the Vmax was an equivalent H+ flux of 15.6 ± 6.3 pmole cm−2 s−1. In Fig. 3, ATPγS supported 16.0 ± 1.3 pmole cm−2 s−1. The agreement between the predicted and observed values is not unreasonable for two reasons. First, we obtained the 36Cl data during continuous dialysis but were forced to halt dialysis for the pHi experiments (Fig. 1). Second, the computed equivalent H+ flux is proportional to the computed buffering power. In fact, if cytosolic components—which have a relatively low natural buffering power (Boron and De Weer 1976b)—diluted the DF, the true closed-system buffering power would have been somewhat less than the computed buffering power of the DF, leading to an overestimate of H+ flux. Note, however, that any error in buffering power (i.e., scaling of the computed equivalent H+ flux) would not affect the computed Km for ATP.
An earlier study with ATPγS (Boron et al. 1988) demonstrated that the squid axon’s Na+-driven Cl–HCO3 exchanger does not use ATP as fuel. One potential mechanism by which low [ATP]i could inhibit Na+-driven Cl–HCO3 exchange would be if, by catalyzing the reaction 2 ADP → ATP + AMP, adenylate kinase would release AMP that would activate AMP kinase, which in turn would block various energy-depleting processes (for review, see Young et al. 2005)—presumably including the Na+-driven Cl–HCO3 exchanger. However, it is not clear how ATPγS could relieve this inhibition except by inhibiting either of the two aforementioned kinases.
One goal of the present study was to rule out one of the remaining hypotheses, namely, that ATP (or ATPγS) promotes transport by either (1) binding to an ATP-binding site or (2) phosphorylating the transporter (or an essential activator). If ATP (or ATPγS) worked via an ATP-binding site, then it would be reasonable to expect that AMP-PNP or AMP-PCP should also bind to the activating site and support transport. Indeed, others have reported that AMP-PNP binds to the cystic fibrosis transmembrane conductance regulator (CFTR) (Hwang et al. 1994; Gunderson and Kopito 1994) and that AMP-PCP inhibits a Cl− channel in zymogen granules (Thevenod et al. 1994). However, we found that neither of these two nonhydrolyzable ATP analogues could support transport, making the binding hypothesis unlikely. Moreover, the cloned squid Na+-driven Cl–HCO3 exchanger sqNDCBE (Virkki et al. 2003) does not have a consensus nucleotide-binding site.
Regarding the phosphorylation hypothesis, it is well established that ATPγS leads to phosphatase-resistant protein phosphorylation (Cassel and Glaser 1982). Other investigators, in studies on membrane proteins, have exploited the stable phosphorylation produced by ATPγS (Wu et al. 2001). Moreover, sqNDCBE (Virkki et al. 2003), on either its cytoplasmic N or C terminus, has multiple protein kinase A and protein kinase C consensus phosphorylation sites, though we do not know whether the phosphorylation state of any of these affects transporter function. We suggest that the most likely hypothesis is that ATPγS similarly leads to the stable phosphorylation of either the Na+-driven Cl–HCO3 exchanger protein or an essential activator.
Two intriguing observations were that, in the presence of a background [ATP]i of ~230 µM, 8 mM AMP-PNP caused a significant decrease in JH, whereas 1 mM ADPβS caused a significant increase (Fig. 4). In principle, AMP-PNP could have inhibited transport by competing with ATP for binding to a hypothetical ATP-binding site. However, because AMP-PNP failed to support transport in the nominal absence of ATP (Fig. 3), we think it is more likely that AMP-PNP inhibited a kinase. In principle, ADPβS could have stimulated transport by binding to a hypothetical ATP-binding site. However, because ADPβS (like AMP-PNP) failed to support transport in the nominal absence of ATP (Fig. 3), we think it is more likely that ADPβS somehow promoted phosphorylation. For example, the axon could have converted ADPβS via adenylate kinase to ATPβS. The γ phosphate of ATPβS (compared to that of ATPγS) is structurally more similar to the γ phosphate of ATP and, thus, might have a higher affinity for the kinase.
This work was supported by National Institutes of Health grants NS18400 and NS11946. We thank Mr. Mike Hernandez for assistance in performing the ATP assays and Mr. Duncan Wong for technical assistance.
Bruce A. Davis, Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, CT 06520, USA.
Emilia M. Hogan, Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, CT 06520, USA.
John M. Russell, Department of Biology, Syracuse University, Syracuse, NY 13244, USA.
Walter F. Boron, Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, CT 06520, USA.