Am J Physiol Cell Physiol 307: C791–C813, 2014. First published June 25, 2014; doi:10.1152/ajpcell.00051.2014.
Evidence from simultaneous intracellular- and surface-pH transients that carbonic anhydrase II enhances CO2 fluxes across Xenopus oocyte plasma membranes Raif Musa-Aziz,1,3,4 Rossana Occhipinti,1 and Walter F. Boron1,2,3 1
Department of Physiology and Biophysics, Case Western Reserve University School of Medicine, Cleveland, Ohio; Department of Medicine, Case Western Reserve University School of Medicine, Cleveland, Ohio; 3Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut; and 4Department of Physiology and Biophysics, University of Sao Paulo, Institute of Biomedical Sciences, Sao Paulo, Brazil 2
Submitted 12 February 2014; accepted in final form 19 June 2014
Musa-Aziz R, Occhipinti R, Boron WF. Evidence from simultaneous intracellular- and surface-pH transients that carbonic anhydrase II enhances CO2 fluxes across Xenopus oocyte plasma membranes. Am J Physiol Cell Physiol 307: C791–C813, 2014. First published June 25, 2014; doi:10.1152/ajpcell.00051.2014.—The ␣-carbonic anhydrases (CAs) are zinc-containing enzymes that catalyze the interconversion of CO2 and HCO⫺ 3 . Here, we focus on human CA II (CA II), a ubiquitous cytoplasmic enzyme. In the second paper in this series, we examine CA IV at the extracellular surface. After microinjecting recombinant CA II in a Tris solution (or just Tris) into oocytes, we expose oocytes to 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 while using microelectrodes to monitor intracellular pH (pHi) and surface pH (pHS). CO2 influx causes the familiar sustained pHi fall as well as a transient pHS rise; CO2 efflux does the opposite. Both during CO2 addition and removal, CA II increases the magnitudes of the maximal rate of pHi change, (dpHi/dt)max, and the maximal change in pHS, ⌬pHS. Preincubating oocytes with the inhibitor ethoxzolamide eliminates the effects of CA II. Compared with pHS, pHi begins to change only after a delay of ⬃9 s and its relaxation has a larger (i.e., slower) time constant (pHi ⬎ pHS). Simultaneous measurements with two pHi electrodes, one superficial and one deep, suggest that impalement depth contributes to pHi delay and higher pHi. Using higher ⫺ ⫺ CO2/HCO⫺ 3 levels, i.e., 5%/33 mM HCO3 or 10%/66 mM HCO3 , increases (dpHi/dt)max and ⌬pHS, though not in proportion to the increase in [CO2]. A reaction-diffusion mathematical model (described in the third paper in this series) accounts for the above general features and supports the conclusion that cytosolic CA— consuming entering CO2 or replenishing exiting CO2—increases CO2 fluxes across the cell membrane. ion-sensitive microelectrodes; ethoxzolamide; electrode depth; electrophysiology; mathematical modeling
␣-CARBONIC ANHYDRASES (CAs) are a family of zinc-containing enzymes that catalyze the interconversion of CO2 and HCO⫺ 3 . Of the 15 human CAs (32, 37, 68), 12 are known to be active (54) in catalyzing the reaction CO2 ⫹ H2O º HCO⫺ 3 ⫹ H⫹, thereby bypassing the slow CO2-hydration/dehydration reactions involving H2CO3. Three CAs—CA VIII (34, 64), CA X (50), and CA XI (23)—lack at least one of the three conserved histidine residues that coordinate Zn2⫹ in other CAs. Because Zn2⫹ is required for carbonic-anhydrase activity, and because the three His residues are required for highaffinity binding of Zn2⫹, it is reasonable to suggest that CA VIII, X, and XI are not catalytically active. Indeed, Sjöblom et THE
Address for reprint requests and other correspondence: R. Musa Aziz, Dept. of Physiology and Biophysics, Univ. of Sao Paulo, Av Prof Lineu Prestes 1524, Sao Paulo, SP 05508-000 Brazil (e-mail: [email protected]). http://www.ajpcell.org
al. (63) restored the CA activity of CA VIII by making two simultaneous point mutations: converting an Arg residue to the His that is conserved in catalytically active ␣-CAs, and converting a Glu residue to a conserved Gln. Although CA VIII, X, and XI do not have clear physiological roles, the catalytically active CAs are major players in a variety of physiological processes that involve CO2 and HCO⫺ 3 transport across cell membranes. These processes include CO2 carriage by erythrocytes (42), HCO⫺ 3 transport across epithelia (11, 21, 36, 53, 54), fluid transport across epithelia (2, 18, 35, 40, 53, 62), transient changes in extracellular pH in the brain (12), and the speed of certain intracellular pH transients (58, 71). For a particular animal species, the enzymatically active ␣-CAs differ in catalytic activity, affinity for CO2, inhibitor sensitivity, and expression pattern (32, 54, 65). Deficiency of CA II is associated with osteopetrosis, renal-tubular acidosis, and cerebral calcification (65). Moreover, CA blockers are used clinically to treat glaucoma (2, 6, 39, 60), metabolic and respiratory alkalosis (3– 6), and acute mountain sickness (4, 6, 55, 69, 74), and are also weak diuretics (4 – 6). In 1973, Gutknecht and Tosteson (30) explored factors that influence the diffusion of salicylic acid across an artificial lipid bilayer. They concluded that the reaction HA º H⫹ ⫹ A⫺ (where A⫺ is the salicylate anion) in the unstirred layers near the membrane, as well as the presence of non-salicylate buffers, plays a major role in enhancing HA flux across the membrane. They also predicted that non-CO2/HCO⫺ 3 buffers would accelerate the flux of CO2. However, in 1977, Gutknecht et al. (29) reported that non-CO2/HCO⫺ 3 buffers (phosphate, HEPES, Tris) enhance CO2 fluxes across lipid bilayers—measured using 14C-labeled CO2— only in the presence of mobile CA. The purpose of the present study, summarized in this paper and its two companions (46, 49), is to elucidate the influence of CAs and non-CO2/HCO⫺ 3 buffers on CO2 fluxes across the plasma membranes of living cells.1 In 1984, De Hemptinne and Huguenin (31) used a miniaturized electrode, with a glass sensor, to monitor the pH at the extracellular surface (pHS) of rat soleus muscle as they applied CO2/HCO⫺ 3 . They observed a transient rise in pHS that, as they pointed out, presumably reflected the influx of CO2, which would promote the following reaction at the extracellular surface of the cell: H⫹ ⫹ HCO⫺ 3 ¡ CO2 ⫹ H2O. We and our collaborators used an adaptation of this pHS approach to study the role of aquaporins 1
This article is the topic of an Editorial Focus by Eric Delpire (18a).
0363-6143/14 Copyright © 2014 the American Physiological Society
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and rhesus proteins as CO2 channels (14, 22, 24 –26, 45). In the present study, we chose to use this adapted pHS approach, as well as simultaneous measurements of intracellular pH (pHi), to assess the CO2 fluxes across the membranes of Xenopus oocytes. In this first paper, we examine the effect of microinjecting oocytes with the soluble enzyme CA II. We chose to work with CA II because it is ubiquitous and has the highest enzymatic activity of all CAs (reviewed in ref. 54). In the second paper (46), we examine the effect of expressing CA IV (GPI-linked to the extracellular cell surface), with and without CA II in the bulk extracellular fluid (BECF). Early exploratory work by Nakhoul et al. (48) on the heterologous expression of CA IV in oocytes motivated portions of this project. In the third paper (49), we extend the three-dimensional reactiondiffusion model of Somersalo et al. (66) to assist in the interpretation of our pHi and pHS data in the other two papers. This model, an an extension of the single-compartment approach developed by Boron and De Weer (9), treats the oocyte as a spherical symmetric cell, with simultaneous reaction and diffusion processes occurring in both the extracellular unconvected fluid (EUF, an extension of the unstirred-layer concept) and the intracellular fluid (ICF). The model accounts for the slow equilibration CO2 ⫹ H2O º H2CO3, effective acceleration of this reaction by CA II and CA IV, and competing equilibria from a multitude of non-CO2/HCO⫺ 3 buffers in both EUF and ICF. Our basic experimental protocol was to expose an oocyte— importantly, an oocyte not expressing any HCO⫺ 3 transporters—to a solution containing 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 (Fig. 1). The CO2 influx causes the familiar fall in pHi (for review, see ref. 58), which intracellular CA ought to accelerate, as first observed by Thomas (71). The influx of CO2 should also reduce [CO2]S. The depleted CO2 at the cell surface is replenished by 1) diffusion from the BECF and 2) the reaction ⫹ HCO⫺ 3 ⫹ H ¡ CO2 ⫹ H2O at the cell surface. Because this
reaction consumes H⫹, pHS rises, as observed by De Hemptinne and Huguenin (31) and by Saarikoski and Kaila (59). The reaction also depletes HCO⫺ 3 at the cell surface, and we would expect that the diffusion of H⫹ and HCO⫺ 3 from the BECF would replenish the depleted H⫹ and HCO⫺ 3 at the cell surface. If an extracellular buffer, such as HEPES, is present, it would undergo the reaction H-HEPES ¡ H⫹ ⫹ HEPES⫺ at the cell surface and thereby reduce the magnitude of the pHS increase. However, according to the experiments of Gutknecht et al. (29), this HEPES reaction should substantially raise [CO2]S (and thus enhance the influx of CO2) in the presence of mobile extracellular CA.2 Over time, as [CO2]i approaches [CO2]S, and as [CO2]S approaches [CO2]BECF, all of these fluxes and reactions slow and the system reaches equilibrium. In this first paper, we find that applying 1.5% CO2/10 mM HCO⫺ 3 (at a fixed pH of 7.50) does indeed cause pHS to rise transiently and then decay with an exponential time course. pHi also falls with an exponential time course, but more slowly, reflecting the depth of the penetration of the pHi electrode into the cell as well as the asymmetry of [CO2]S as BECF flows past the oocyte. Injecting CA II into the oocyte not only accelerates the decline in pHi but also increases the height and the rate of decay of the pHS spike. The CA II inhibitor ethoxzolamide (EZA) blocks all of the effects of CA II. Our pHS data imply that, by ⫹ speeding the conversion of incoming CO2 to HCO⫺ 3 ⫹ H , intracellular CA II keeps [CO2]i relatively low and thus enhances the gradient driving CO2 influx. In turn, this increased CO2 influx enhances the fall in [CO2]S, and thus accentuates the pHS spike. A mathematical model, which is discussed in the third paper in this series (49), supports this hypothesis. 2 In the companion paper (46), we tested these predictions concerning the extracellular buffer.
EUF
EUF Fig. 1. Model of an oocyte exposed to CO2/HCO⫺ 3 . The main part of the figure illustrates the diffusion and reaction events as CO2 enters an oocyte. The black arrows with sharp heads indicate solute diffusion in the extracellular unconvected fluid (EUF) and intracellular fluid. The black arrows with dull heads indicate reactions. The left inset is a schematic top view of the oocyte in the chamber, as seen through the microscope. The orange arrows indicate the direction of convective flow of the bulk extracellular fluid (BECF). The right inset is a schematic view of the oocyte along the axis of convective flow, looking downstream. The darker half of the oocyte, oriented upward, is the animal pole. CA II, carbonic anhydrase II; pHi, intracellular pH; pHS, surface pH.
A
CO2
[CO2]S
H+
H 2O
HCO3–
CO2
CO2 H2O B
pHi
CA II
CA IV
[HCO3– ] HCO3–
H+
HCO3–
BECF
BECF Top View
Direction of solution flow
Downstream View Vm
Vm pHi
pHi pHS
pHS
Meridian
(20 μm tip)
AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
Equator Chamber bottom
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Preparation of Xenopus Oocytes We isolated stage V-VI oocytes from Xenopus laevis using standard techniques (27). Briefly, we anesthetized animals in 0.2% MS222 (ethyl 3-aminobenzoate methanesulfonate, Sigma-Aldrich, St. Louis, MO) and removed ovarian lobes, which we placed in a 0-Ca solution (98 mM NaCl, 2 mM KCl, 1 mM MgCl2, 5 mM HEPES, pH 7.50). After washing the oocytes 5⫻ in the 0-Ca solution, we dissociated and defolliculated them by enzymatic digestion, using 2 mg/ml type IA collagenase (Sigma-Aldrich) in the 0-Ca solution. We selected stage V-VI oocytes and kept them at 18°C in sterile-filtered OR3 medium, each 2 liters of which contained one pack of powdered Leibovitz L-15 media with L-glutamine (GIBCO-BRL), 100 ml of 10,000 U/ml penicillin, 10,000 U/ml streptomycin solution (SigmaAldrich), and 5 mM HEPES titrated to pH 7.50, and osmolality adjusted to 195 mosmol/kgH2O. One day after their isolation, we injected the oocytes with 50 nl of H2O as a control for the injection of cRNA encoding CA IV in the accompanying paper (46). Three days after this H2O injection, we injected the oocytes with 50 nl of fluid, either Tris buffer (50 mM titrated to pH 7.5 with HCl) or Tris buffer containing 6 ng/nl of recombinant human CA II protein (i.e., 300 ng/oocyte ⬵ 10 pmol, for a final [CA II]i of ⬃30 M, ⬃50% higher than the level in red blood cells). Dr. Peter Piermarini kindly prepared the CA II, as described previously (38, 52), using an approach similar to that described by others (70), and assessing protein purity by polyacrylamide gel electrophoresis (PAGE) and MALDI mass spectrometry (SynPep, Dublin, CA). After the Tris or CA II injection, we incubated the oocytes overnight at 18°C in the sterile-filtered OR3 medium described above. The protocols for housing and handling of Xenopus laevis were approved by the Institutional Animal Care and Use Committee of Case Western Reserve and Yale Universities. Solutions Table 1 summarizes the composition of the solutions. The ND96 ⫺ solution is nominally CO2/HCO⫺ 3 free. The CO2/HCO3 solutions were bubbled vigorously with the appropriate gas mixture (balance O2) for ⬃30 min. We dissolved the CA II inhibitor EZA (Sigma-Aldrich) in 0.05 N NaOH, to prepare a stock solution with a final concentration of 50 mM. We achieved a final EZA concentration of 400 M by diluting this stock 1:125 in ND96, and adjusted the pH to 7.50 with 5 N HCl. Electrophysiological Measurements Figure 8B in Ref. 44 provides a detailed view of the arrangement of the chamber, oocyte, and electrodes. Chamber. We placed an oocyte in a plastic perfusion chamber with a channel 3 mm wide ⫻ 30 mm long. A glass coverslip formed the
bottom of the chamber. Solutions were placed in 140-ml plastic syringes (Sherwood Medical, St. Louis, MO) and delivered to the chamber using syringe pumps (Harvard Apparatus, South Natick, MA). Solutions were carried to the chamber via Tygon tubing (Ryan Herco Products, Burbank, CA; Formulation R3603-3; OD 4.8 mm /ID 1.6 mm) and flowed from one end of the 30-mm channel to the other. In initial experiments for the data set summarized below in Fig. 3, the solution flowed at 2 ml/min, and in later experiments, 3 ml/min. As noted in RESULTS, it does not appear that this change had a significant effect. For all other data sets in this paper, solutions flowed at 3 ml/min. We switched among solutions with pneumatically operated valves (Clippard Instrument Laboratory, Cincinnati, OH). All experiments were performed at room temperature (⬃22°C), and in all experiments, the oocyte was initially superfused with the ND96 solution. Measurement of intracellular pH. In a typical experiment, we impaled the oocyte (with the dark animal pole facing upward) with two microelectrodes (Fig. 1, left and right insets), one for sensing membrane potential (Vm, amplified by a model OC-725 two-electrode Oocyte Clamp, Warner Instruments, Hamden, CT) and the other for sensing pHi (amplified by a model FD223 high-impedance electrometer, World Precision Instruments, Sarasota, FL). We fabricated and used the electrodes as described previously (56, 57, 61). We used a horizontal microelectrode puller (model P-97, Sutter Instrument, Novato, CA) to produce microelectrodes from thin-walled borosilicate glass (part no. G200TF-4, 2.0 mm OD ⫻1.56 mm ID, Warner Instruments). We filled the Vm electrodes with 3 M KCl; these had resistances of ⬃0.6 M⍀. The pHi electrodes were identical but we filled them with a liquid, pH-sensitive membrane (Hydrogen Ionophore I, mixture B, Fluka Chemical, Ronkonkoma, NY). The extracellular reference for the Vm electrode was the virtual ground created by the Warner OC-725 Oocyte Clamp. The ISENSE connection was attached to a microelectrode holder (model 64-1010, Warner Instruments), which contained a Ag/AgCl half-cell that served as a bridge to a broken-tipped glass microelectrode filled with 3 M KCl (⬃0.1 M⍀), and positioned so that its tip was close to the oocyte. The IOUT connection was attached to a platinum wire that rested in a reservoir at the end of the chamber’s channel (i.e., downstream from the oocyte). The circuit ground of the OC-725 (i.e., the virtual ground of the BECF or bath), and indeed the circuit grounds of all electronic components, was connected to a brass plate (i.e., “system ground”), which was in turn connected to earth. We obtained the voltage due to pHi by electronically subtracting the signal of the Vm electrode from that of the pHi electrode. We obtained Vm by electronically subtracting the system ground of the OC-725 from the signal of the Vm electrode. The device that performed the subtractions (Yale University Subtraction Amplifier, V3.1) also appropriately scaled the voltages for the inputs of an analog-todigital converter within a Windows-based computer. We simultaneously
Table 1. Solutions 1.5% CO2/10 mM HCO⫺ 3
5% CO2/33 mM HCO⫺ 3
10% CO2/66 mM HCO⫺ 3
Component
ND96 Sol. 1
1 Sol. 2
HEPES 5 Sol. 3
25 Sol. 4
1 Sol. 5
HEPES 5 Sol. 6
25 Sol. 7
1 Sol. 8
HEPES 5 Sol. 9
25 Sol. 10
NaCl, mM* KCl, mM MgCl2, mM CaCl2, mM HEPES, mM* HCO⫺ 3 , mM pH Osmolality, mosmol/kgH2O
96 2 1 1.8 5 0 7.5 ⬵195
88 2 1 1.8 1 10 7.5 ⬵195
86 2 1 1.8 5 10 7.5 ⬵195
76 2 1 1.8 25 10 7.5 ⬵195
65 2 1 1.8 1 33 7.5 ⬵195
63 2 1 1.8 5 33 7.5 ⬵195
53 2 1 1.8 25 33 7.5 ⬵195
32 2 1 1.8 1 66 7.5 ⬵195
30 2 1 1.8 5 66 7.5 ⬵195
20 2 1 1.8 25 66 7.5 ⬵195
*We titrated HEPES free acid (pK 7.5) to pH 7.5 with NaOH (so that, theoretically, we have incremented H-HEPES, HEPES⫺, and Na⫹ concentrations by equal amounts) and then secondarily adjusted [NaCl], as indicated in this table, to achieve a final osmolality of ⬃195 mosmol/kgH2O. AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
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acquired all electrical data, initially, once every 2,000 ms, later once every 1,000 ms, and finally once every 500 ms, and analyzed it using software written in-house. As noted in the DISCUSSION, these differences do not appear to affect our data. We obtained the slope of the intracellular pH electrodes by calibrating pH standards at pH 6.0 and 8.0 (model SB 104-1 and SB 112-1, respectively, Fisher Scientific, Fair Lawn, NJ). An additional single-point calibration was obtained in the ND96 solution (pH 7.50) in the bath before impaling the oocyte. Oocytes had a spontaneous Vm at least as negative as ⫺40 mV. Measurement of surface pH. In addition to the electrodes described above, in a typical experiment we measured pHS using a liquid-ionexchange pH electrode (Fig. 1, left and right insets) with an outer tip diameter of 20 m (amplified by a FD223 electrometer, WPI). We pulled these pipettes from standard-wall borosilicate tubing (part no. G200F-4, 2.0 mm OD ⫻1.16 mm ID, Warner Instruments) and used a microforge to break off and fire polish the tips as one would for a giant-patch pipette (33). During experiments, we attach the pHS electrode to an ultra-fine computer-controlled micromanipulator with digital position display (model MPC-200 system, Sutter Instrument). A typical experiment begins with the pHS electrode tip in the bath—in the “home” position, ⬃300 m from the oocyte surface—to calibrate the electrode at pH 7.50 in ND96 solution (Table 1). After that, we move the flat tip of the pHS electrode to the oocyte’s surface—the “zero” position—near the equator (i.e., between the animal and vegetal poles, halfway between top and bottom), and ⬃5° behind the meridian perpendicular to the axis of flow (i.e., slightly in the “shadow” of the flowing extracellular solution), as illustrated in the left inset of Fig. 1. Finally, we further advance the electrode tip ⬃40 m until we observe a slight dimple in the membrane. The creation of this small dimple produces a microenvironment between the pHS electrode tip and the oocyte membrane and has two effects: 1) maximizing the pHS change and 2) ensuring a consistent and reproducible electrode placement, in the radial direction, on the oocyte membrane for each experiment, and thus minimizing variability from experiment to experiment. During the experiment, we periodically withdraw the electrode to its “home” position for recalibration in the flowing BECF solution at pH 7.50. In the figures, we indicate these movements of the pHS electrode in the radial direction by “Surface” and “BECF” in the step chart. The external reference for the pHS electrode was a very long, broken-tipped (⬃10 m ID) glass micropipette that we pulled from the above from thin-walled borosilicate tubing on a rotating micropipette puller (model 51.511, Stoelting, Chicago, IL), filled with 3 M KCl, and bridged with a calomel half-cell to the input of a model 750 electrometer (WPI). pHS was obtained by subtracting the calomel signal from the pHS signal. As for the pHi electrodes, we obtained the slope of the pHS electrodes by calibrating pH standards at pH 6.0 and 8.0. In preliminary experiments, we found that the slope of the electrodes was the same in ND96 vs. CO2/HCO⫺ 3 . Dual measurement of intracellular pH. In some experiments, we replaced the above pHS microelectrode with a second sharp pHi microelectrode (designated electrode no. 2), which we moved using the same Sutter 200 micromanipulator that otherwise carried the pHS electrode. Rate of bath solution change. In mock experiments, we used pHi and pHS electrodes (sampling 1 per 500 ms) to monitor the pH of the bath (flowing at 3 ml/min) as we switched from a pH-7.5 to a pH-8.0 solution. We found that the time constant () for the reported time course of pH was 1.76 ⫾ 0.13 s (n ⫽ 3) for pHi and 1.70 ⫾ 0.35 s (n ⫽ 4) for pHS. Because these figures include the response time of the electrodes, ⬃1.7 s is the maximum for switching the solution in the chamber. Analysis of pH Data Intracellular pH. Applying extracellular CO2/HCO⫺ 3 causes a fall in the measured pHi that at first begins slowly (presumably reflecting the finite time required for [CO2]BECF to rise to its maximal value, the depth of penetration of the electrode into the cell, and the asymmetry
of [CO2]S over the oocyte surface) and then accelerates to the maximal rate of decline—(dpHi/dt)max. The rate of pHi decline then slows and gradually falls to zero. For example, in CA II experiments such as that in Fig. 2A below (summarized in Fig. 3 below), the fall in pHi accelerated for an average of 9 ⫾ 1 s (n ⫽ 16) before reaching the period of steepest decline. By inspection, we determined the time
A
“CA II” surface electrode position:
Surface
BECF
1.5% CO2 10 mM HCO3– 7.7
ΔpHS
pHS
7.5
7.3
pHi
pH
ΔpHi
7.1
5 min 6.9
−0.0021
B
+0.0013
(dpHi/dt)max
6.7
“Tris”
1.5% CO2 10 mM HCO3– 7.7
pHS
7.5
7.3
pH pHi
7.1
6.9
5 min
−0.0009 +0.0006
6.7
Fig. 2. Representative experiments showing effects of CA II on pHi and pHS changes evoked by application and removal of CO2/HCO⫺ 3 . A: oocytes injected with recombinant human CA II dissolved in Tris buffer. B: oocytes injected only with Tris buffer. Both in “CA II” or “Tris” oocytes, the pHi trace is represented by the red lower record, and pHS, by the green upper record. At the indicated times, we switched the extracellular solution from ND96 to 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 (Table 1) and then back again. In this example, the extracellular solution flowed at 2 ml/min and the sampling rate was 1 per 1,000 ms. The vertical gray bands represent periods during which the pHS electrode was withdrawn to the BECF for calibration. At other times the pHS electrode was dimpling ⬃40 m into the oocyte surface. The gray, dashed vertical lines represent the times that the computer switched the valves to initiate a change of solutions. The left and right blue, dashed vertical lines (see RESULTS) represent the initiation of the pHS and pHi transients, respectively. The dashed black lines through the initial portions of the pHi records for CO2 application and removal represent best linear fits for maximal rates of pHi change (negative or positive direction). The downward vertical arrows near the pHi records represent the CO2-induced changes in steady-state pHi. The upward and downward arrows near the pHS records represent maximal changes in pHS (positive or negative direction).
AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
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“CA II”
A
“Tris” P=10-4
0.003
P=0.009
(dpHi/dt)max (pHi·s-1)
0.002 0.001
I ⫽ ⫺
0.000
(16)
(16)
−0.001 −0.002
P=0.4 −0.003
P=0.03
CO2 addition
CO2 removal
B 0.0 −0.1
ΔpHi −0.2 −0.3
(16)
(16) P=0.97
C
P
Evidence from simultaneous intracellular- and surface-pH transients that carbonic anhydrase II enhances CO2 fluxes across Xenopus oocyte plasma membranes Raif Musa-Aziz,1,3,4 Rossana Occhipinti,1 and Walter F. Boron1,2,3 1
Department of Physiology and Biophysics, Case Western Reserve University School of Medicine, Cleveland, Ohio; Department of Medicine, Case Western Reserve University School of Medicine, Cleveland, Ohio; 3Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut; and 4Department of Physiology and Biophysics, University of Sao Paulo, Institute of Biomedical Sciences, Sao Paulo, Brazil 2
Submitted 12 February 2014; accepted in final form 19 June 2014
Musa-Aziz R, Occhipinti R, Boron WF. Evidence from simultaneous intracellular- and surface-pH transients that carbonic anhydrase II enhances CO2 fluxes across Xenopus oocyte plasma membranes. Am J Physiol Cell Physiol 307: C791–C813, 2014. First published June 25, 2014; doi:10.1152/ajpcell.00051.2014.—The ␣-carbonic anhydrases (CAs) are zinc-containing enzymes that catalyze the interconversion of CO2 and HCO⫺ 3 . Here, we focus on human CA II (CA II), a ubiquitous cytoplasmic enzyme. In the second paper in this series, we examine CA IV at the extracellular surface. After microinjecting recombinant CA II in a Tris solution (or just Tris) into oocytes, we expose oocytes to 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 while using microelectrodes to monitor intracellular pH (pHi) and surface pH (pHS). CO2 influx causes the familiar sustained pHi fall as well as a transient pHS rise; CO2 efflux does the opposite. Both during CO2 addition and removal, CA II increases the magnitudes of the maximal rate of pHi change, (dpHi/dt)max, and the maximal change in pHS, ⌬pHS. Preincubating oocytes with the inhibitor ethoxzolamide eliminates the effects of CA II. Compared with pHS, pHi begins to change only after a delay of ⬃9 s and its relaxation has a larger (i.e., slower) time constant (pHi ⬎ pHS). Simultaneous measurements with two pHi electrodes, one superficial and one deep, suggest that impalement depth contributes to pHi delay and higher pHi. Using higher ⫺ ⫺ CO2/HCO⫺ 3 levels, i.e., 5%/33 mM HCO3 or 10%/66 mM HCO3 , increases (dpHi/dt)max and ⌬pHS, though not in proportion to the increase in [CO2]. A reaction-diffusion mathematical model (described in the third paper in this series) accounts for the above general features and supports the conclusion that cytosolic CA— consuming entering CO2 or replenishing exiting CO2—increases CO2 fluxes across the cell membrane. ion-sensitive microelectrodes; ethoxzolamide; electrode depth; electrophysiology; mathematical modeling
␣-CARBONIC ANHYDRASES (CAs) are a family of zinc-containing enzymes that catalyze the interconversion of CO2 and HCO⫺ 3 . Of the 15 human CAs (32, 37, 68), 12 are known to be active (54) in catalyzing the reaction CO2 ⫹ H2O º HCO⫺ 3 ⫹ H⫹, thereby bypassing the slow CO2-hydration/dehydration reactions involving H2CO3. Three CAs—CA VIII (34, 64), CA X (50), and CA XI (23)—lack at least one of the three conserved histidine residues that coordinate Zn2⫹ in other CAs. Because Zn2⫹ is required for carbonic-anhydrase activity, and because the three His residues are required for highaffinity binding of Zn2⫹, it is reasonable to suggest that CA VIII, X, and XI are not catalytically active. Indeed, Sjöblom et THE
Address for reprint requests and other correspondence: R. Musa Aziz, Dept. of Physiology and Biophysics, Univ. of Sao Paulo, Av Prof Lineu Prestes 1524, Sao Paulo, SP 05508-000 Brazil (e-mail: [email protected]). http://www.ajpcell.org
al. (63) restored the CA activity of CA VIII by making two simultaneous point mutations: converting an Arg residue to the His that is conserved in catalytically active ␣-CAs, and converting a Glu residue to a conserved Gln. Although CA VIII, X, and XI do not have clear physiological roles, the catalytically active CAs are major players in a variety of physiological processes that involve CO2 and HCO⫺ 3 transport across cell membranes. These processes include CO2 carriage by erythrocytes (42), HCO⫺ 3 transport across epithelia (11, 21, 36, 53, 54), fluid transport across epithelia (2, 18, 35, 40, 53, 62), transient changes in extracellular pH in the brain (12), and the speed of certain intracellular pH transients (58, 71). For a particular animal species, the enzymatically active ␣-CAs differ in catalytic activity, affinity for CO2, inhibitor sensitivity, and expression pattern (32, 54, 65). Deficiency of CA II is associated with osteopetrosis, renal-tubular acidosis, and cerebral calcification (65). Moreover, CA blockers are used clinically to treat glaucoma (2, 6, 39, 60), metabolic and respiratory alkalosis (3– 6), and acute mountain sickness (4, 6, 55, 69, 74), and are also weak diuretics (4 – 6). In 1973, Gutknecht and Tosteson (30) explored factors that influence the diffusion of salicylic acid across an artificial lipid bilayer. They concluded that the reaction HA º H⫹ ⫹ A⫺ (where A⫺ is the salicylate anion) in the unstirred layers near the membrane, as well as the presence of non-salicylate buffers, plays a major role in enhancing HA flux across the membrane. They also predicted that non-CO2/HCO⫺ 3 buffers would accelerate the flux of CO2. However, in 1977, Gutknecht et al. (29) reported that non-CO2/HCO⫺ 3 buffers (phosphate, HEPES, Tris) enhance CO2 fluxes across lipid bilayers—measured using 14C-labeled CO2— only in the presence of mobile CA. The purpose of the present study, summarized in this paper and its two companions (46, 49), is to elucidate the influence of CAs and non-CO2/HCO⫺ 3 buffers on CO2 fluxes across the plasma membranes of living cells.1 In 1984, De Hemptinne and Huguenin (31) used a miniaturized electrode, with a glass sensor, to monitor the pH at the extracellular surface (pHS) of rat soleus muscle as they applied CO2/HCO⫺ 3 . They observed a transient rise in pHS that, as they pointed out, presumably reflected the influx of CO2, which would promote the following reaction at the extracellular surface of the cell: H⫹ ⫹ HCO⫺ 3 ¡ CO2 ⫹ H2O. We and our collaborators used an adaptation of this pHS approach to study the role of aquaporins 1
This article is the topic of an Editorial Focus by Eric Delpire (18a).
0363-6143/14 Copyright © 2014 the American Physiological Society
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and rhesus proteins as CO2 channels (14, 22, 24 –26, 45). In the present study, we chose to use this adapted pHS approach, as well as simultaneous measurements of intracellular pH (pHi), to assess the CO2 fluxes across the membranes of Xenopus oocytes. In this first paper, we examine the effect of microinjecting oocytes with the soluble enzyme CA II. We chose to work with CA II because it is ubiquitous and has the highest enzymatic activity of all CAs (reviewed in ref. 54). In the second paper (46), we examine the effect of expressing CA IV (GPI-linked to the extracellular cell surface), with and without CA II in the bulk extracellular fluid (BECF). Early exploratory work by Nakhoul et al. (48) on the heterologous expression of CA IV in oocytes motivated portions of this project. In the third paper (49), we extend the three-dimensional reactiondiffusion model of Somersalo et al. (66) to assist in the interpretation of our pHi and pHS data in the other two papers. This model, an an extension of the single-compartment approach developed by Boron and De Weer (9), treats the oocyte as a spherical symmetric cell, with simultaneous reaction and diffusion processes occurring in both the extracellular unconvected fluid (EUF, an extension of the unstirred-layer concept) and the intracellular fluid (ICF). The model accounts for the slow equilibration CO2 ⫹ H2O º H2CO3, effective acceleration of this reaction by CA II and CA IV, and competing equilibria from a multitude of non-CO2/HCO⫺ 3 buffers in both EUF and ICF. Our basic experimental protocol was to expose an oocyte— importantly, an oocyte not expressing any HCO⫺ 3 transporters—to a solution containing 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 (Fig. 1). The CO2 influx causes the familiar fall in pHi (for review, see ref. 58), which intracellular CA ought to accelerate, as first observed by Thomas (71). The influx of CO2 should also reduce [CO2]S. The depleted CO2 at the cell surface is replenished by 1) diffusion from the BECF and 2) the reaction ⫹ HCO⫺ 3 ⫹ H ¡ CO2 ⫹ H2O at the cell surface. Because this
reaction consumes H⫹, pHS rises, as observed by De Hemptinne and Huguenin (31) and by Saarikoski and Kaila (59). The reaction also depletes HCO⫺ 3 at the cell surface, and we would expect that the diffusion of H⫹ and HCO⫺ 3 from the BECF would replenish the depleted H⫹ and HCO⫺ 3 at the cell surface. If an extracellular buffer, such as HEPES, is present, it would undergo the reaction H-HEPES ¡ H⫹ ⫹ HEPES⫺ at the cell surface and thereby reduce the magnitude of the pHS increase. However, according to the experiments of Gutknecht et al. (29), this HEPES reaction should substantially raise [CO2]S (and thus enhance the influx of CO2) in the presence of mobile extracellular CA.2 Over time, as [CO2]i approaches [CO2]S, and as [CO2]S approaches [CO2]BECF, all of these fluxes and reactions slow and the system reaches equilibrium. In this first paper, we find that applying 1.5% CO2/10 mM HCO⫺ 3 (at a fixed pH of 7.50) does indeed cause pHS to rise transiently and then decay with an exponential time course. pHi also falls with an exponential time course, but more slowly, reflecting the depth of the penetration of the pHi electrode into the cell as well as the asymmetry of [CO2]S as BECF flows past the oocyte. Injecting CA II into the oocyte not only accelerates the decline in pHi but also increases the height and the rate of decay of the pHS spike. The CA II inhibitor ethoxzolamide (EZA) blocks all of the effects of CA II. Our pHS data imply that, by ⫹ speeding the conversion of incoming CO2 to HCO⫺ 3 ⫹ H , intracellular CA II keeps [CO2]i relatively low and thus enhances the gradient driving CO2 influx. In turn, this increased CO2 influx enhances the fall in [CO2]S, and thus accentuates the pHS spike. A mathematical model, which is discussed in the third paper in this series (49), supports this hypothesis. 2 In the companion paper (46), we tested these predictions concerning the extracellular buffer.
EUF
EUF Fig. 1. Model of an oocyte exposed to CO2/HCO⫺ 3 . The main part of the figure illustrates the diffusion and reaction events as CO2 enters an oocyte. The black arrows with sharp heads indicate solute diffusion in the extracellular unconvected fluid (EUF) and intracellular fluid. The black arrows with dull heads indicate reactions. The left inset is a schematic top view of the oocyte in the chamber, as seen through the microscope. The orange arrows indicate the direction of convective flow of the bulk extracellular fluid (BECF). The right inset is a schematic view of the oocyte along the axis of convective flow, looking downstream. The darker half of the oocyte, oriented upward, is the animal pole. CA II, carbonic anhydrase II; pHi, intracellular pH; pHS, surface pH.
A
CO2
[CO2]S
H+
H 2O
HCO3–
CO2
CO2 H2O B
pHi
CA II
CA IV
[HCO3– ] HCO3–
H+
HCO3–
BECF
BECF Top View
Direction of solution flow
Downstream View Vm
Vm pHi
pHi pHS
pHS
Meridian
(20 μm tip)
AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
Equator Chamber bottom
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CA II ENHANCES CO2 FLUXES MATERIALS AND METHODS
Preparation of Xenopus Oocytes We isolated stage V-VI oocytes from Xenopus laevis using standard techniques (27). Briefly, we anesthetized animals in 0.2% MS222 (ethyl 3-aminobenzoate methanesulfonate, Sigma-Aldrich, St. Louis, MO) and removed ovarian lobes, which we placed in a 0-Ca solution (98 mM NaCl, 2 mM KCl, 1 mM MgCl2, 5 mM HEPES, pH 7.50). After washing the oocytes 5⫻ in the 0-Ca solution, we dissociated and defolliculated them by enzymatic digestion, using 2 mg/ml type IA collagenase (Sigma-Aldrich) in the 0-Ca solution. We selected stage V-VI oocytes and kept them at 18°C in sterile-filtered OR3 medium, each 2 liters of which contained one pack of powdered Leibovitz L-15 media with L-glutamine (GIBCO-BRL), 100 ml of 10,000 U/ml penicillin, 10,000 U/ml streptomycin solution (SigmaAldrich), and 5 mM HEPES titrated to pH 7.50, and osmolality adjusted to 195 mosmol/kgH2O. One day after their isolation, we injected the oocytes with 50 nl of H2O as a control for the injection of cRNA encoding CA IV in the accompanying paper (46). Three days after this H2O injection, we injected the oocytes with 50 nl of fluid, either Tris buffer (50 mM titrated to pH 7.5 with HCl) or Tris buffer containing 6 ng/nl of recombinant human CA II protein (i.e., 300 ng/oocyte ⬵ 10 pmol, for a final [CA II]i of ⬃30 M, ⬃50% higher than the level in red blood cells). Dr. Peter Piermarini kindly prepared the CA II, as described previously (38, 52), using an approach similar to that described by others (70), and assessing protein purity by polyacrylamide gel electrophoresis (PAGE) and MALDI mass spectrometry (SynPep, Dublin, CA). After the Tris or CA II injection, we incubated the oocytes overnight at 18°C in the sterile-filtered OR3 medium described above. The protocols for housing and handling of Xenopus laevis were approved by the Institutional Animal Care and Use Committee of Case Western Reserve and Yale Universities. Solutions Table 1 summarizes the composition of the solutions. The ND96 ⫺ solution is nominally CO2/HCO⫺ 3 free. The CO2/HCO3 solutions were bubbled vigorously with the appropriate gas mixture (balance O2) for ⬃30 min. We dissolved the CA II inhibitor EZA (Sigma-Aldrich) in 0.05 N NaOH, to prepare a stock solution with a final concentration of 50 mM. We achieved a final EZA concentration of 400 M by diluting this stock 1:125 in ND96, and adjusted the pH to 7.50 with 5 N HCl. Electrophysiological Measurements Figure 8B in Ref. 44 provides a detailed view of the arrangement of the chamber, oocyte, and electrodes. Chamber. We placed an oocyte in a plastic perfusion chamber with a channel 3 mm wide ⫻ 30 mm long. A glass coverslip formed the
bottom of the chamber. Solutions were placed in 140-ml plastic syringes (Sherwood Medical, St. Louis, MO) and delivered to the chamber using syringe pumps (Harvard Apparatus, South Natick, MA). Solutions were carried to the chamber via Tygon tubing (Ryan Herco Products, Burbank, CA; Formulation R3603-3; OD 4.8 mm /ID 1.6 mm) and flowed from one end of the 30-mm channel to the other. In initial experiments for the data set summarized below in Fig. 3, the solution flowed at 2 ml/min, and in later experiments, 3 ml/min. As noted in RESULTS, it does not appear that this change had a significant effect. For all other data sets in this paper, solutions flowed at 3 ml/min. We switched among solutions with pneumatically operated valves (Clippard Instrument Laboratory, Cincinnati, OH). All experiments were performed at room temperature (⬃22°C), and in all experiments, the oocyte was initially superfused with the ND96 solution. Measurement of intracellular pH. In a typical experiment, we impaled the oocyte (with the dark animal pole facing upward) with two microelectrodes (Fig. 1, left and right insets), one for sensing membrane potential (Vm, amplified by a model OC-725 two-electrode Oocyte Clamp, Warner Instruments, Hamden, CT) and the other for sensing pHi (amplified by a model FD223 high-impedance electrometer, World Precision Instruments, Sarasota, FL). We fabricated and used the electrodes as described previously (56, 57, 61). We used a horizontal microelectrode puller (model P-97, Sutter Instrument, Novato, CA) to produce microelectrodes from thin-walled borosilicate glass (part no. G200TF-4, 2.0 mm OD ⫻1.56 mm ID, Warner Instruments). We filled the Vm electrodes with 3 M KCl; these had resistances of ⬃0.6 M⍀. The pHi electrodes were identical but we filled them with a liquid, pH-sensitive membrane (Hydrogen Ionophore I, mixture B, Fluka Chemical, Ronkonkoma, NY). The extracellular reference for the Vm electrode was the virtual ground created by the Warner OC-725 Oocyte Clamp. The ISENSE connection was attached to a microelectrode holder (model 64-1010, Warner Instruments), which contained a Ag/AgCl half-cell that served as a bridge to a broken-tipped glass microelectrode filled with 3 M KCl (⬃0.1 M⍀), and positioned so that its tip was close to the oocyte. The IOUT connection was attached to a platinum wire that rested in a reservoir at the end of the chamber’s channel (i.e., downstream from the oocyte). The circuit ground of the OC-725 (i.e., the virtual ground of the BECF or bath), and indeed the circuit grounds of all electronic components, was connected to a brass plate (i.e., “system ground”), which was in turn connected to earth. We obtained the voltage due to pHi by electronically subtracting the signal of the Vm electrode from that of the pHi electrode. We obtained Vm by electronically subtracting the system ground of the OC-725 from the signal of the Vm electrode. The device that performed the subtractions (Yale University Subtraction Amplifier, V3.1) also appropriately scaled the voltages for the inputs of an analog-todigital converter within a Windows-based computer. We simultaneously
Table 1. Solutions 1.5% CO2/10 mM HCO⫺ 3
5% CO2/33 mM HCO⫺ 3
10% CO2/66 mM HCO⫺ 3
Component
ND96 Sol. 1
1 Sol. 2
HEPES 5 Sol. 3
25 Sol. 4
1 Sol. 5
HEPES 5 Sol. 6
25 Sol. 7
1 Sol. 8
HEPES 5 Sol. 9
25 Sol. 10
NaCl, mM* KCl, mM MgCl2, mM CaCl2, mM HEPES, mM* HCO⫺ 3 , mM pH Osmolality, mosmol/kgH2O
96 2 1 1.8 5 0 7.5 ⬵195
88 2 1 1.8 1 10 7.5 ⬵195
86 2 1 1.8 5 10 7.5 ⬵195
76 2 1 1.8 25 10 7.5 ⬵195
65 2 1 1.8 1 33 7.5 ⬵195
63 2 1 1.8 5 33 7.5 ⬵195
53 2 1 1.8 25 33 7.5 ⬵195
32 2 1 1.8 1 66 7.5 ⬵195
30 2 1 1.8 5 66 7.5 ⬵195
20 2 1 1.8 25 66 7.5 ⬵195
*We titrated HEPES free acid (pK 7.5) to pH 7.5 with NaOH (so that, theoretically, we have incremented H-HEPES, HEPES⫺, and Na⫹ concentrations by equal amounts) and then secondarily adjusted [NaCl], as indicated in this table, to achieve a final osmolality of ⬃195 mosmol/kgH2O. AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
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acquired all electrical data, initially, once every 2,000 ms, later once every 1,000 ms, and finally once every 500 ms, and analyzed it using software written in-house. As noted in the DISCUSSION, these differences do not appear to affect our data. We obtained the slope of the intracellular pH electrodes by calibrating pH standards at pH 6.0 and 8.0 (model SB 104-1 and SB 112-1, respectively, Fisher Scientific, Fair Lawn, NJ). An additional single-point calibration was obtained in the ND96 solution (pH 7.50) in the bath before impaling the oocyte. Oocytes had a spontaneous Vm at least as negative as ⫺40 mV. Measurement of surface pH. In addition to the electrodes described above, in a typical experiment we measured pHS using a liquid-ionexchange pH electrode (Fig. 1, left and right insets) with an outer tip diameter of 20 m (amplified by a FD223 electrometer, WPI). We pulled these pipettes from standard-wall borosilicate tubing (part no. G200F-4, 2.0 mm OD ⫻1.16 mm ID, Warner Instruments) and used a microforge to break off and fire polish the tips as one would for a giant-patch pipette (33). During experiments, we attach the pHS electrode to an ultra-fine computer-controlled micromanipulator with digital position display (model MPC-200 system, Sutter Instrument). A typical experiment begins with the pHS electrode tip in the bath—in the “home” position, ⬃300 m from the oocyte surface—to calibrate the electrode at pH 7.50 in ND96 solution (Table 1). After that, we move the flat tip of the pHS electrode to the oocyte’s surface—the “zero” position—near the equator (i.e., between the animal and vegetal poles, halfway between top and bottom), and ⬃5° behind the meridian perpendicular to the axis of flow (i.e., slightly in the “shadow” of the flowing extracellular solution), as illustrated in the left inset of Fig. 1. Finally, we further advance the electrode tip ⬃40 m until we observe a slight dimple in the membrane. The creation of this small dimple produces a microenvironment between the pHS electrode tip and the oocyte membrane and has two effects: 1) maximizing the pHS change and 2) ensuring a consistent and reproducible electrode placement, in the radial direction, on the oocyte membrane for each experiment, and thus minimizing variability from experiment to experiment. During the experiment, we periodically withdraw the electrode to its “home” position for recalibration in the flowing BECF solution at pH 7.50. In the figures, we indicate these movements of the pHS electrode in the radial direction by “Surface” and “BECF” in the step chart. The external reference for the pHS electrode was a very long, broken-tipped (⬃10 m ID) glass micropipette that we pulled from the above from thin-walled borosilicate tubing on a rotating micropipette puller (model 51.511, Stoelting, Chicago, IL), filled with 3 M KCl, and bridged with a calomel half-cell to the input of a model 750 electrometer (WPI). pHS was obtained by subtracting the calomel signal from the pHS signal. As for the pHi electrodes, we obtained the slope of the pHS electrodes by calibrating pH standards at pH 6.0 and 8.0. In preliminary experiments, we found that the slope of the electrodes was the same in ND96 vs. CO2/HCO⫺ 3 . Dual measurement of intracellular pH. In some experiments, we replaced the above pHS microelectrode with a second sharp pHi microelectrode (designated electrode no. 2), which we moved using the same Sutter 200 micromanipulator that otherwise carried the pHS electrode. Rate of bath solution change. In mock experiments, we used pHi and pHS electrodes (sampling 1 per 500 ms) to monitor the pH of the bath (flowing at 3 ml/min) as we switched from a pH-7.5 to a pH-8.0 solution. We found that the time constant () for the reported time course of pH was 1.76 ⫾ 0.13 s (n ⫽ 3) for pHi and 1.70 ⫾ 0.35 s (n ⫽ 4) for pHS. Because these figures include the response time of the electrodes, ⬃1.7 s is the maximum for switching the solution in the chamber. Analysis of pH Data Intracellular pH. Applying extracellular CO2/HCO⫺ 3 causes a fall in the measured pHi that at first begins slowly (presumably reflecting the finite time required for [CO2]BECF to rise to its maximal value, the depth of penetration of the electrode into the cell, and the asymmetry
of [CO2]S over the oocyte surface) and then accelerates to the maximal rate of decline—(dpHi/dt)max. The rate of pHi decline then slows and gradually falls to zero. For example, in CA II experiments such as that in Fig. 2A below (summarized in Fig. 3 below), the fall in pHi accelerated for an average of 9 ⫾ 1 s (n ⫽ 16) before reaching the period of steepest decline. By inspection, we determined the time
A
“CA II” surface electrode position:
Surface
BECF
1.5% CO2 10 mM HCO3– 7.7
ΔpHS
pHS
7.5
7.3
pHi
pH
ΔpHi
7.1
5 min 6.9
−0.0021
B
+0.0013
(dpHi/dt)max
6.7
“Tris”
1.5% CO2 10 mM HCO3– 7.7
pHS
7.5
7.3
pH pHi
7.1
6.9
5 min
−0.0009 +0.0006
6.7
Fig. 2. Representative experiments showing effects of CA II on pHi and pHS changes evoked by application and removal of CO2/HCO⫺ 3 . A: oocytes injected with recombinant human CA II dissolved in Tris buffer. B: oocytes injected only with Tris buffer. Both in “CA II” or “Tris” oocytes, the pHi trace is represented by the red lower record, and pHS, by the green upper record. At the indicated times, we switched the extracellular solution from ND96 to 1.5% CO2/10 mM HCO⫺ 3 /pH 7.50 (Table 1) and then back again. In this example, the extracellular solution flowed at 2 ml/min and the sampling rate was 1 per 1,000 ms. The vertical gray bands represent periods during which the pHS electrode was withdrawn to the BECF for calibration. At other times the pHS electrode was dimpling ⬃40 m into the oocyte surface. The gray, dashed vertical lines represent the times that the computer switched the valves to initiate a change of solutions. The left and right blue, dashed vertical lines (see RESULTS) represent the initiation of the pHS and pHi transients, respectively. The dashed black lines through the initial portions of the pHi records for CO2 application and removal represent best linear fits for maximal rates of pHi change (negative or positive direction). The downward vertical arrows near the pHi records represent the CO2-induced changes in steady-state pHi. The upward and downward arrows near the pHS records represent maximal changes in pHS (positive or negative direction).
AJP-Cell Physiol • doi:10.1152/ajpcell.00051.2014 • www.ajpcell.org
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“CA II”
A
“Tris” P=10-4
0.003
P=0.009
(dpHi/dt)max (pHi·s-1)
0.002 0.001
I ⫽ ⫺
0.000
(16)
(16)
−0.001 −0.002
P=0.4 −0.003
P=0.03
CO2 addition
CO2 removal
B 0.0 −0.1
ΔpHi −0.2 −0.3
(16)
(16) P=0.97
C
P
