Jiang Q , Mak D , Devidas S , Schwiebert EM , Bragin A , Zhang Y , Skach WR , Guggino WB , Foskett JK , Engelhardt JF .

DK48977 NIDDK NIH HHS , DK49136 NIDDK NIH HHS , HL47122 NHLBI NIH HHS , P50 DK049136 NIDDK NIH HHS , R01 HL047122 NHLBI NIH HHS

	Figure 2. Absence of ATP efflux after activation of Ca+-activated Cl− conductance in oocytes. To assess whether membrane potential changes due to opening of Cl− channels in oocytes could nonspecifically facilitate ATP efflux, the rates of ATP efflux in cAMP/forskolin stimulated CFTR cRNA- injected oocytes, and ionomycin-treated water-injected oocytes were compared. A demonstrates the mean (± SEM) relative light units produced from oocytes derived from two frogs after exposure to the following sequence of buffers: (1) 140 mM Na-gluconate/HPBR; (2) 140 mM Na-gluconate/HPBR + (200 μM 8-cpt-cAMP, 2.5 μM forskolin, or 10 μM ionomycin); and (3) 140 mM NaCl/HPBR + (200 μM 8-cpt-cAMP, 2.5 μM forskolin, or 10 μM ionomycin). Buffer changes are denoted by arrows, and the treatment conditions are indicated in the legend. B depicts the mean (± SEM) rate of ATP efflux for each buffer condition as calculated from the slope of relative light units vs. time (for the 100-s reading) for each individual oocyte. ATP concentration curve standards were used to calibrate rates in terms of pmoles/min (1 pmole ATP = 90 light units). All oocytes analyzed were included in these results. To demonstrate that ionomycin has no effect on the activity of luciferin–luciferase, we compared standard curves for ATP in the presence of buffers and agonists used for the above study (C). Ionomycin standard curves were measured in Na-gluconate buffers.
	Figure 3. Comparison of CFTR-modulated ATP release between wild-type, TMD1, and Δ259-M265V CFTR cRNA– injected Xenopus oocytes. Results in A demonstrate the mean (± SEM) relative light units produced by CFTR cRNA–injected oocytes exposed to the following order of buffers: (1) 140 mM Na-gluconate/HPBR; (2) 140 mM Na-gluconate/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin; and (3) 140 mM NaCl/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin. Buffer changes are denoted by arrows, and cRNA injected is indicated in the legend. B depicts the mean (± SEM) rate of ATP efflux for each buffer condition as calculated from the slope of relative light units vs. time (for the 100-s reading) for each individual oocyte. C depicts the mean (± SEM) Wc chloride conductance values representing the total and delta conductance after adding forskolin/IBMX to the bathing solution (corrected for water-injected background). D gives the mean I/V relationships for each construct in the presence and absence of the cAMP agonists forskolin/IBMX. An equal number (N) of wt CFTR, TMD1, and Δ259-M265V cRNA-injected oocytes were assessed for ATP efflux within each of two batches of oocytes. All oocytes analyzed were included in these results derived from two independent frogs.
	Figure 4. CFTR-modulated ATP release is influenced by extracellular halides. A depicts the mean (± SEM) relative light units produced by CFTR cRNA-injected oocytes (N = 25) from five frogs. Oocytes were exposed to the following order of buffers: (1) 140 mM Na-gluconate/HPBR; (2) 140 mM Na-gluconate/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin; (3) 140 mM NaCl/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin; (4) 140 mM NaBr/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin; and (5) 140 mM NaCl/HPBR, 200 μM 8-cpt-cAMP, 2.5 μM forskolin. Buffer changes are denoted by arrows. Only oocytes that demonstrated release in the presence of extracellular Cl− (arrow 3) were pooled for this analysis. The mean (± SEM) rates of ATP efflux calculated from the rates for each individual oocyte were 3.3 ± 1.5 pmoles ATP/min (buffer 3), 1.0 ± 0.5 pmoles ATP/min (buffer 4), and 4.0 ± 1.7 pmoles ATP/min (buffer 5). B depicts the affects of various halides and anions on the activity of luciferin–luciferase using standard curves for ATP (no differences were seen in Cl and Br-containing buffers). The halide selectivity of CFTR-modulated ATP release in SOS and HPBR was compared in C. These studies evaluated ATP efflux in wtCFTR cRNA-injected oocytes using the specified order of buffers as shown in the figure (details of buffer composition are described in Materials and Methods). Results depict the mean (± SEM, N = 6) relative light units produced in each buffer with mean (± SEM) rates of ATP efflux shown above each line. Calculated rates of ATP efflux are the mean of rates from individual oocytes in pmoles ATP/min. In this analysis, oocytes that demonstrated no ATP efflux under all of the buffer conditions were not included in these calculations that attempted to compare Cl to Br dependence of ATP release.
	Figure 5. Extracellular Cl− concentration determines the relative activity of CFTR-modulated ATP release. Two independent batches of wtCFTR cRNA- injected Xenopus oocytes were analyzed for the effects of extracellular Cl− concentration on the activation of ATP efflux. An HPBR solution was used throughout these experiments with alterations in the final NaCl concentration facilitated by exchange of an equal molar amount of Na-gluconate with NaCl. The total concentration of NaCl and Na-gluconate was equal to 140 mM throughout these experiments with a constant osmolarity. A demonstrates the mean (± SEM) relative light units produced by CFTR-expressing oocytes for which the extracellular Cl− concentration was incrementally increased by adding NaCl-containing buffers with a constant concentration of luciferin–luciferase enzymes. Between seven and eleven oocytes were evaluated at each Cl− concentration from two independent frogs. At the end of the experiment, Cl− was replaced with Br− to confirm the halide selectivity of the oocytes analyzed. Oocytes that did not demonstrate ATP efflux above water-injected controls (data not shown) were not included in this data set. B depicts the mean (± SEM) rates of ATP efflux from cumulative data of four independent batches of CFTR cRNA-injected oocytes as a function of Cl− concentration. The rate of ATP efflux in the presence of extracellular Br− is also included for reference. The concentration of extracellular Cl− necessary for activation of CFTR-modulated ATP efflux was ∼115 mM (arrow).
	Figure 6. Electrophysiological properties of CFTR mutants using Wc two-electrode voltage clamp measurements. Mutants of CFTR that alter charged arginine residues within the channel pore including R334W, R347E, and R347P were used. A demonstrates the amount of [35S]methionine-labeled CFTR immunoprecipitated with anti-hCFTR antibodies from oocytes injected with mutant and wild-type CFTR cRNAs. The arrow depicts unglycosylated CFTR protein produced 3 h after injection. The Wc I/V relationships for wtCFTR, R347P, R347E, R334W cRNA, and water-injected oocytes are given in B. Arrows denote the I/V relations before forskolin/IBMX activation in the presence of extracellular Cl− (marked baseline) and after adding forskolin/ IBMX in the presence of extracellular Cl− (Cl) and bromide (Br). Stimulation of CFTR was performed by perfusion of the oocyte with appropriate buffers containing 100 μM IBMX and 10 μM forskolin. ND96 buffer was used throughout the experiment where NaBr replaced NaCl for halide selectivity measurements. The order of buffer perfusion was as follows: (a) ND96 (NaCl); (b) ND96 (NaCl) + forskolin/IBMX; and (c) ND96 (NaBr) + forskolin/IBMX. These results depict average I/V relationships from N experiments for wtCFTR (N = 5), R347E (N = 5), R347P (N = 5), R334W (N = 8), and water (N = 7)-injected oocytes from at least two independent batches of oocytes.
	Figure 7. Halide dependence of CFTR-modulated ATP release is mediated by a chloride sensor within the channel pore. Results in A demonstrate the mean (± SEM) relative light units produced after activation with 8-cpt-cAMP/forskolin and changes in the extracellular halide concentration. The number of oocytes (N) compiled for each mutant are given with the number of independent experiments for wtCFTR = 3, R347P = 2, R334E = 3, R334W = 2, and water = 3. Although all mutants were not always evaluated at one time, CFTR and water-injected controls were always performed alongside of mutants to confirm the integrity of ATP responses in the particular batch of oocytes. B depicts the mean (± SEM) rate of ATP efflux during the last three buffer conditions as calculated from the slope of relative light units vs. time (for the 100-s reading) for each individual oocyte. The halide selectivity of CFTR-modulated ATP release is compared with the halide permeability (PBr/PCl) and conductance (GBr/GCl) ratios for each of the mutants in C. Results depict the mean (± SEM) for each construct as determined from the individual I/V curves (the number of oocytes analyzed are shown in Table II).
	Figure 8. Chloride sensor model for CFTR activation of ATP release. The proposed mechanism for CFTR-modulated ATP release in Xenopus oocytes must fit several criteria. First, extracellular Cl− and cAMP agonist stimulation are necessary but not sufficient to activate CFTR-modulated ATP release. Second, the fact that not all batches of oocytes that demonstrated CFTR-modulated Cl− conductance were capable of ATP efflux, suggests that a cofactor(s) (blue) is needed for CFTR-modulated ATP release (CFTR is yellow). The dependency of CFTR-modulated ATP release on the external Cl− concentration and the effects of mutagenesis of putative pore residues in CFTR suggest that Cl− may bind within the channel pore to activate structural changes in CFTR necessary for the appropriate interactions with a cofactor(s)- dependent ATP release pathway. This model also incorporates the altered halide selectivity of CFTR mediated conductance (Cl = Br) and ATP efflux (Cl > Br) as dependent on the affinity for halide binding at the chloride sensor within the channel pore.

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