Domain-domain associations in cystic fibrosis transmembrane conductance regulator

Wenlan Wang1, Zhaoping He1, Thomas J. O'Shaughnessy1, John Rux2, and William W. Reenstra3

1 Alfred I. duPont Hospital for Children, Wilmington, Delaware 19803; 2 Wistar Institute and 3 Institute for Human Gene Therapy, University of Pennsylvania, Philadelphia, Pennsylvania 19104


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cystic fibrosis is caused by mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) gene. CFTR is a chloride channel whose activity requires protein kinase A-dependent phosphorylation of an intracellular regulatory domain (R-domain) and ATP hydrolysis at the nucleotide-binding domains (NBDs). To identify potential sites of domain-domain interaction within CFTR, we expressed, purified, and refolded histidine (His)- and glutathione-S-transferase (GST)-tagged cytoplasmic domains of CFTR. ATP-binding to his-NBD1 and his-NBD2 was demonstrated by measuring tryptophan fluorescence quenching. Tryptic digestion of in vitro phosphorylated his-NBD1-R and in situ phosphorylated CFTR generated the same phosphopeptides. An interaction between NBD1-R and NBD2 was assayed by tryptophan fluorescence quenching. Binding among all pairwise combinations of R-domain, NBD1, and NBD2 was demonstrated with an overlay assay. To identify specific sites of interaction between domains of CFTR, an overlay assay was used to probe an overlapping peptide library spanning all intracellular regions of CFTR with his-NBD1, his-NBD2, and GST-R-domain. By mapping peptides from NBD1 and NBD2 that bound to other intracellular domains onto crystal structures for HisP, MalK, and Rad50, probable sites of interaction between NBD1 and NBD2 were identified. Our data support a model where NBDs form dimers with the ATP-binding sites at the domain-domain interface.

tryptophan fluorescence; peptide array; molecular structure; peptide library; phosphorylation


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

CYSTIC FIBROSIS (CF) is a genetic disease caused by mutations in the gene encoding the cystic fibrosis transmembrane conductance regulator (CFTR), an epithelial chloride channel (35, 37). CFTR is a member of the ATP-binding cassette (ABC) transporter gene superfamily (14). More than 1,000 ABC transporters are known, and they are found in both prokaryotic and eukaryotic cells where they carry out the unidirectional ATP-dependent transport of a wide variety of molecules. All known ABC transporters have two membrane-spanning domains (MSDs), usually composed of six transmembrane helices, and two nucleotide-binding domains (NBDs) (14). These domains can be expressed in a single subunit, as in CFTR and multidrug resistance protein (MDR), or in multiple subunits, as in the bacterial maltose and histidine transporters. ABC transporters like CFTR and MDR have intracellular NH2- and COOH-terminals and four intracellular loops, L1 to L4, between transmembrane helices 2 and 3, 4 and 5, 8 and 9, and 10 and 11, respectively. CFTR is unique among ABC family members because it is the only family member to contain an additional intracellular domain, the regulatory or R-domain. CFTR is also the only ABC family member that is known to function as an ion channel (37). The R-domain contains multiple protein kinase A (PKA)-dependent phosphorylation sites that are involved in the regulation of CFTR channel activity (35). The regulation of CFTR channel activity involves a two-step process; phosphorylation of the R-domain, most likely on multiple sites, is required so that ATP hydrolysis by the NBDs can regulate CFTR gating (1, 11, 34, 36, 46). Mutations in the R-domain, most notably at phosphorylation sites, alter ATP-dependent CFTR gating (46). In addition, the CF-causing Delta F508 mutation, located in NBD1, has been shown to alter in situ phosphorylation (18). The fact that alterations in one domain can modulate the function of a second domain indicates that functionally important domain-domain interactions must occur (11). The primary goal of the study was to identify the sites of domain-domain interaction.

A number of studies have suggested that specific domain-domain associations are required for optimal CFTR activity. An in vitro association between NBD1-R and NBD2 constructs has been demonstrated by fluorescence quenching and exclusion chromatography (26). Interactions between the NH2-and COOH-terminals with other domains of CFTR have been postulated on the basis of CFTR channel activity (29, 32, 42). However, there is no information on the specific sites of interaction among CFTR domains. In addition, recent electrophysiological and cytochemical studies have suggested that native CFTR may exist as a homodimer (9, 32, 42, 47), but any dimeric form would appear to be relatively unstable because biochemical studies have failed to detect multimeric forms of CFTR (6, 28). Despite the fact that CFTR has been shown by both immunological and yeast two-hybrid studies to interact with a number of membrane-associated proteins (12, 13, 29, 39).

No crystal structures have been generated for any domain of CFTR; however, structures for bacterial NBDs have been reported. These include HisP from the histidine transporter of Salmonella typhimurium and MalK from the maltose transporter of Thermococcus litoralis (7, 17). In addition, a crystal structure for Rad50, a DNA-binding protein with a similar motif to HisP and MalK, has been reported (15). Comparison of the crystal structures for the NBD monomers suggests that the basic fold is conserved. Recently, two additional crystal structures for NBD monomers have been reported; they have the same structure as the NBDs of HisP and MalK (21, 44). However, crystallographic structures for HisP, MalK, and Rad50 dimers vary considerably. Because the relative orientation of the two NBDs should be conserved among all ABC family members (19), we suggest that no more than one, and possibly none, of the observed dimeric structures are related to the structure in native ABC transporters. Although additional crystal structures may resolve this issue, we have taken a biochemical approach. To this end, individual cytoplasmic domains of CFTR were expressed, purified, and refolded. Sites of domain-domain association were determined by in vitro binding to an overlapping peptide library generated from the intracellular regions of CFTR. Peptides within the NBDs that bound cytoplasmic domains were mapped onto the known NBD structures. Predicted sites of domain-domain interaction were compared with published structures for NBD dimers.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Generation of expression constructs. Polymerase chain reaction (PCR) was employed to amplify cDNA fragments of CFTR from a pcDNA vector (Invitrogen) containing CFTR (obtained from Dr. W. Skach, Oregon Health Sciences University). DNA fragments of NBD1 (nucleotides 1249-1899; amino acids 373-589), NBD2 (nucleotides 3583-4560; amino acids 1151-1476), and NBD1-R (nucleotides 1249-2709; amino acids 373-859) were cloned into pProEx HT (Qiagen) with an NH2-terminal his-x6 tag. Because of its poor expression as a His-tagged protein, R-domain (nucleotides 1891-2709; amino acids 589-830) was cloned into pGEX-5X (Amersham Pharmacia Biotech) with glutathione-S-transferase (GST) fused to its NH2 terminus. The correct recombinants were identified by restriction mapping and sequencing.

Expression, purification, and refolding. CFTR recombinants were expressed in Escherichia coli BL21codon-plus (Strategene). His-tagged CFTR domains were purified from inclusion bodies in 8 M urea by Ni-affinity chromatography according to the manufacturer's instruction (Qiagen). GST-R protein was purified on glutathione-Sepharose 4B (10). Protein purity was analyzed by SDS-PAGE, and protein concentrations were determined with bicinchoninic acid (40). GST-R domain was renatured by dialyzing overnight against phosphate-buffered saline (PBS, pH 7.2). His-tagged proteins were renatured by diluting 10-fold into binding buffer (50 mM Tris, pH 7.5, 0.15 M NaCl, 1 mM EDTA, 0.5% NP-40, 0.5 mM dithiothreitol, and 0.1 mg/ml BSA) for peptide binding assays or PBS for fluorescence studies.

Fluorescence studies. Fluorescence measurements were performed at room temperature using a Perkin-Elmer LS50B spectrofluorometer. Samples were excited at 295 nm and emission spectra were recorded at 345 nm. Excitation and emission bandwidths were 5.0 nm, and spectra were corrected for background fluorescence. For measurements of ATP-dependent fluorescence quenching, ATP was added to domains in binding buffer and the percent change in fluorescence was calculated at each ATP concentration. For the assay of domain-domain association, separate samples of a single domain and two domains were generated for each experimental condition.

Phosphorylation and two-dimensional peptide mapping. In situ CFTR labeling was performed as previously described (33). Briefly, NIH-3T3 cells stably expressing human CFTR (NIH-CFTR) were incubated with 32Pi for 2 h before stimulation with 10 µM forskolin for 2 min. Cells were lysed in 4°C lysis buffer (100 mM NaCl, 50 mM NaF, 0.1% SDS, 1% Na-deoxycholate, 1% Triton X-100, 1 mM EDTA, 1 mM EGTA, 0.1 mM phenylmethylsulfonyl fluoride, 0.1 mg/mL aprotinin, 1 mM orthovanadate, and 50 mM Tris · HCl, pH 7.5) and lysate-cleared by centrifugation. CFTR was immunoprecipitated with a COOH-terminal CFTR antibody (R&D) and protein A beads (Calbiochem) and was then purified by SDS-PAGE. For in vitro labeling, CFTR was immunoprecipitated from NIH-CFTR cells as described, with the exception that cells were not exposed to 32Pi or forskolin. Immunoprecipitated CFTR was suspended in 50 µl of kinase buffer (50 mM Tris, 10 mM MgCl2, and 100 µg/ml BSA, pH 7.5) and phosphorylated with 2 units of PKA catalytic subunit (Sigma) and 10 µCi [gamma -32P]ATP. CFTR was resolved by SDS-PAGE. For in vitro labeling of NBD1-R, purified protein (1-2 µg) was placed in kinase buffer plus 0.8 M urea and phosphorylated with 2 units of PKA and 10 µCi [gamma -32P]ATP. Phosphorylated NBD1-R was separated from the reaction mixture by SDS-PAGE. In all cases, phosphorylated products were visualized with a Storm 860 PhosphorImager.

Two-dimensional phosphopeptide mapping of phosphorylated CFTR and phosphorylated NBD1-R was performed as previously described (2, 45). Phosphorylated CFTR and NBD-R were extracted from the gel and TCA was precipitated. Protein was reacted with trypsin overnight and lyophilized before peptide separation by two-dimensional (2-D) peptide mapping electrophoresis at pH 8.9 and ascending chromatography in butanol-pyridine-acetic acid-water (15:10:3:12 vol:vol:vol:vol). Phosphopeptides were visualized with a Storm 860 PhosphorImager (Molecular Dynamics) and identified by comparison with published (5, 33) and unpublished (Dr. J. Cohn) phosphopeptide maps of CFTR.

Peptide binding assay. Peptide walking was used to identify sites of domain-domain interaction (20). One hundred fifty-seven overlapping 20-mer peptides, spanning all cytoplasmic regions of CFTR, were synthesized by the multipin synthesis method (41) and were purity-analyzed on HPLC (Chiron). Peptides overlapped by 13 residues. All peptides were acetylated at the NH2 terminus and amidated at the COOH terminus. Peptides were dissolved in 100% DMSO to a concentration of 2 mM and stored frozen at -70 °C. For binding assays, peptides in DMSO were diluted with distilled water (1:10), and then 10 µg of each peptide were blotted onto polyvinylide difluoride membrane (Bio-Rad). The membranes were air-dried, blocked with 5% milk in PBST (PBS-0.5% Tween), and then incubated with 10 µg/ml of a purified CFTR domain overnight in binding buffer. Bound proteins were detected with monoclonal antibodies against GST or His tags (Upstate Biotechnology) or with a polyclonal antibody against NBD2 (generated against the expressed domain by Covance Research Products). Signals were visualized by enhanced chemiluminescence (Amersham Pharmacia Biotech).


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Expression and characterization of CFTR domains. His-tagged expression vectors for NBD1, NBD2, and the NBD1-R-domain of CFTR were constructed. In addition, we expressed a GST-tagged R-domain because of poor expression of His-tagged R-domain. These proteins, his-NBD1 (amino acids 373-589), his-NBD2 (amino acids 1151-1476), his-NBD1-R (amino acids 373-859), and GST-R (amino acids 589-830), were expressed in E. coli and purified by affinity chromatography. As shown in Fig. 1, the purified domains contained >95% of the intended product. His-NBD1, his-NBD2, and GST-R migrated at the expected molecular masses of 24, 36, and 52 kDa, respectively, whereas his-NBD1-R migrated at 60 kDa as opposed to the predicted molecular mass of 53 kDa. In all cases, the identity of the expressed domain was confirmed by Western blotting with the antibodies against individual domains (data not shown).


View larger version (97K):
[in this window]
[in a new window]
 
Fig. 1.   Expression and purification of cystic fibrosis transmembrane conductance regulator (CFTR) domains. Purified His-tagged nucleotide-binding domains his-NBD1 (lane 1), his-NBD2 (lane 2), his-NBD1-R (lane 3), and glutathione-S-transferase-tagged R-domain GST-R (lane 4) were run on SDS-PAGE (10%), and proteins were stained with Coomassie blue. Molecular mass markers are indicated at left.

Purified domains were refolded as described in MATERIALS AND METHODS. To establish that the purified domains assumed a "native-like" structure, ATP binding to domains containing an NBD was assayed by measuring ATP-dependent changes in tryptophan fluorescence. All NBD-containing domains bound ATP. ATP binding was concentration dependent (Fig. 2), and disassociation constants for his-NBD1 and his-NBD2 were calculated to be 660 ± 70 and 65 ± 9 µM, respectively. To assess the structural integrity of his-NBD1-R and GST-R, the refolded proteins were phosphorylated in vitro with PKA and [gamma -32P]ATP and digested with trypsin, and phosphopeptides were separated by 2-D peptide mapping. In addition, wild-type CFTR, stably expressed in transfected NIH-3T3 cells, was in situ phosphorylated during stimulation with forskolin and in vitro phosphorylated with PKA and [gamma -32P]ATP after immunoprecipitation. Phosphorylated proteins were digested with trypsin and phosphopeptides were separated by 2-D peptide mapping. As shown in Fig. 3, the same phosphopeptides were generated by in vitro phosphorylation of his-NBD1-R and in situ phosphorylation of CFTR (data for GST-R not shown). However, in vitro phosphorylation of CFTR resulted in a different pattern of phosphorylation (note the absence of phosphorylation on serines 737 and 813). Although the precise locations of peptides in the three maps differed, this was largely due to differences in the relative separation in the horizontal (electrophoretic) and vertical (chromatographic) directions. Significantly, all phosphopeptides from NBD1-R corresponded to a phosphopeptide in the map for in situ phosphorylated CFTR, whereas in vitro phosphorylated CFTR that had been dissolved in detergent and then resuspended in a detergent-free buffer before phosphorylation showed a distinct pattern of phosphorylation. This suggests that NBD1-R has a structure more similar to that of CFTR in situ than that of in vitro phosphorylated CFTR.


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 2.   ATP-dependent quenching of NBD fluorescence. The tryptophan fluorescence (excitation at 295 nm, emission at 345 nm) of 2.0 µM his-NBD1 () or his-NBD2 () in PBS plus 0.8 M urea was assayed as a function of ATP concentration. After subtraction of buffer fluorescence, relative fluorescence was calculated as (FNBD + ATP/FNDB) where FNBD + ATP and FNBD are the fluorescences of NBD in the presence and absence of ATP. Data are presented as means ± SE (n = 3). Lines indicate hyperbolic fits to the data.



View larger version (40K):
[in this window]
[in a new window]
 
Fig. 3.   Tryptic 2-dimensional phosphopeptide maps of CFTR and his-NBD1-R. In vitro phosphorylated CFTR (A), in vitro phosphorylated his-NBD1-R (B), and in situ phosphorylated CFTR (C) were isolated and digested with trypsin as described in MATERIALS AND METHODS. The resulting peptides were separated by electrophoresis in the horizontal direction and ascending chromatography in the vertical direction. Phosphopeptides were located with a Storm 860 PhosphorImager (Molecular Dynamics). Phosphopeptides were identified (site of phosphorylation) by comparison with published (5, 33) and unpublished (Dr. J. Cohn) phosphopeptide maps of CFTR.

To determine whether expressed domains formed domain-domain associations in vitro, changes in tryptophan fluorescence were monitored when expressed his-NBD1-R and his-NBD2 were mixed together. As shown in Fig. 4A, the fluorescence of his-NBD1-R was a linear function of concentration. Based on the assumption that his-NBD2 and his-NBD1-R do not interact, the fluorescence of his-NBD1-R/his-NBD2 mixtures was calculated. These values are indicated by square symbols in Fig. 4A. However, as shown by triangle symbols, the observed fluorescence was significantly less than the predicted value. In Fig. 4A, inset, the percentage decrease in observed fluorescence is plotted as a function of his-NBD1-R concentration. The data were fit to a hyperbolic function with an apparent dissociation constant for his-NBD1-R binding to 1.5 µM his-NBD2 of 0.7 ± 0.3 µM. A weaker association between his-NBD1 and his-NBD2 was also observed (data not shown). Under these experimental conditions, the fluorescence of all expressed domains was linear with respect to concentration, as was the increase in fluorescence from the addition of BSA (data not shown). To determine whether the fluorescence quenching altered ATP binding, we compared the ATP-dependent fluorescence quenching of his-NBD1-R, his NBD2, and a mixture of the two. As shown in Fig. 4B, ATP quenched the fluorescence of both of his-NBD1-R and his-NBD2. However, it caused a significant increase in the fluorescence of a mixture of his-NBD1-R and his-NBD2. Although calculation of the amounts of free and associated domains from the data in Fig. 4A is not warranted, it is clear that both free and associated domains must exist and, because ATP binding to the free domains decreases tryptophan fluorescence, the increase in fluorescence when ATP binds to the associated domains must be greater than that indicated by the change in total fluorescence. We have also used an overlay assay to demonstrate associations between the expressed domains. As shown in Fig. 5A, GST-R, but not GST, bound to his-NBD1 and his-NBD2. An association between his-NBD1 and his-NBD2 was also observed (Fig. 5B).


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 4.   Association of his-NBD2 and his-NBD1-R. A: the tryptophan fluorescence (excitation at 295 nm, emission at 345 nm) of his-NBD1-R alone () and his-NBD1-R in the presence of 1.5 µM his-NBD2 (black-triangle) (Fobs) were determined. Also shown is the calculated fluorescence (Fcal) of a mixture of 1.5 µM his-NBD1-R and the indicated concentration of his-NBD2 () (FNBD1-R + FNBD2-FBUFFER). Data are representative of 3 experiments. Inset, the %decrease in fluorescence [(Fcal - Fobs)/Fcal] is plotted as a function of his-NBD1-R concentration. Line indicates a hyperbolic fit to the data. B: the ATP-dependent change in fluorescence for the addition of 5 mM ATP to 2 µM his-NBD1-R, 1.5 µM his-NBD2, and a mixture of 2 µM his-NBD1-R and 1.5 µM his-NBD2 is shown. Data are presented as means ± SE (n = 3).



View larger version (56K):
[in this window]
[in a new window]
 
Fig. 5.   Overlay assay of domain-domain binding. A: his-NBD1 and his-NBD2 were spotted onto nitrocellulose, overlaid with GST or GST-R and probed with an antibody to GST. B: GST, GST-R, and his-NBD1 are spotted, overlaid with his-NBD2, and probed with an antibody to NBD2. Controls where his-NBD2 was omitted were negative. For A and B, bound domains were visualized by chemiluminescence. Data are representative of 2 assays.

Binding of expressed domains to peptides from CFTR. To identify epitopes that are involved in domain-domain interactions, an overlapping peptide library of 20-mers spanning all intracellular amino acids of CFTR was synthesized. Peptides 1-10, 11-17, 18-25, 97-103, and 105-111 spanned the NH2 terminus and intracellular loops L1-L4, respectively. Peptides 26-96 spanned NBD1 and the R-domain, whereas peptides 112-157 spanned NBD2 and the COOH-terminus. The identity of each peptide is indicated in Fig. 6A. As shown in Fig. 6B, each peptide was assayed for the ability to bind the expressed domains with an overlay assay. Peptides were spotted onto nitrocellulose membranes and incubated with a domain, and bound domain was probed with antibodies to the domain or the fused tag. To identify false positives, the binding of antibodies, or GST plus anti-GST, in the absence of expressed domains was determined. Our anti-His tag antibody bound to none of our peptides, whereas the anti-NBD2 antibody bound to epitopes on three overlapping peptides (135-137) from NBD2 (data not shown). As shown in Fig. 6A, GST was bound by peptides 11, 12, 48, 63, 96, and 106, and more protein was bound to peptide 13. In addition, all three expressed domains bound to peptides 13, 59, 60, 96, and 106, indicated in gray in Fig. 6A; these five interactions were considered false positives and not included in the following analysis. In addition, there are undoubtedly peptides that form associations with a domain in the native structure but do not bind the domain in our assay. These false negatives could occur if the association between peptide and domain were to weak, if the peptide were not retained on the membrane, or if spatial restrictions prevented peptide-domain interaction on the membrane. It is likely that some of the more hydrophilic peptides were lost from the membrane because the percentage of bound peptides and Kyte-Dolittle hydrophobicity (24) were correlated (R2 = 0.91; data not shown). Therefore, it is unlikely that we have identified all sites of domain-domain interaction. Because some peptides were likely to be lost from the membrane, comparisons were made only on the basis of the amount of bound domain and not of the relative strength of association. Despite the uncertainty caused by the potential loss of peptides, the specificity of domain binding can be assessed for any peptide that shows differential binding to the tested domains. For example, data with his-NBD2 and GST-R indicate that peptides 135-137 and 144-148 are retained on the membrane. Because there is no evidence for the binding of his-NBD1 to these peptides, these peptides establish a background for nonspecific binding by NBD1.


View larger version (47K):
[in this window]
[in a new window]
 
Fig. 6.   Binding of GST, R-domain, NBD1, and NBD2 to peptides from the cytoplasmic domains of CFTR. A: the numbered bars indicate the sequence of each of the 157 overlapping peptides. The sequences contained in NBD1, NBD2, and the R-domain constructs are highlighted in red, blue, and yellow, respectively. Peptides in the 12 transmembrane helices are indicated in red. Peptides that bind to NBD1, NBD2, or the R-domain are colored red, blue, or yellow, respectively, with lighter colors indicating less bound domain. Peptides with 2 colors bound both of the indicated domains. Five peptides that bound all 3 domains are shown in gray. B: blots of bound GST, GST-R, his-NBD1, and his-NBD2 are shown. Peptides were blotted onto nitrocellulose membranes, incubated overnight with 1 µg/ml protein, and probed with a polyclonal antibody to the His tag, NBD2, or GST. Bound domains were visualized by chemiluminescence.

As shown in Figure 6B, GST-R bound to peptides 1, 2, and 3 in the NH2 terminus; peptides 14, 16, 105, 109, and 110 in intracellular loops L1 and L4; peptides 46, 48, 49, 53, and 55 in NBD1; and peptides 115, 129, 130, 137, 138, 140, and 144-148 in NBD2 (indicated in yellow in Fig. 6A). His-NBD1 and his-NBD2 had distinct patterns of peptide binding, although they also bound to several peptides in common. His-NBD1 bound to peptides 1, 2, 3, and 10 from the NH2 terminus; peptides 11, 12, and 25 from intracellular loops L1 and L2; peptides 130, 131, 132, 139, and 140 from NBD2; and peptide 48 from NBD1 (indicated in red in Fig. 6A). In contrast, his-NBD2 bound to peptide 20 from intracellular loop L2; peptides 44, 45, 46, and 63 in NBD1; and peptides 131, 132, and 145-148 in NBD2 (indicated in blue in Fig. 6A). Note that peptide 63, although present in our R-domain construct, has recently been shown to be in NBD1 (3). Associations were observed between his-NBD2 and peptides from all four intracellular loops. Two additional points should be made. Whereas R-domain bound to peptides from NBD1 and NBD2, peptides from the R-domain did not bind to any domain. Associations between NBD1 and peptides from NBD1, as well as NBD2 and peptides in NBD2, may reflect a similarity between the structures of NBD1 and NBD2 or self-associations between the NBDs that are present in a dimeric form of CFTR (9, 32, 42, 47). At present, we have no reason to favor either possibility.

Mapping of binding data to known NBD structures. To better understand the relationship of the binding studies to the structure of the NBD1 and NBD2, we have mapped the binding sites in NBD1 and NBD2 onto crystallographic structures of the HisP and MalK monomers, two bacterial NBDs (7, 17). These proteins are overlaid in Fig. 7A. They have sequence identities of only 26%, but comparison of the crystal structures by combinatorial extension (38) gives a root mean square deviation (RMSD) for the alpha -carbons of 2.7 Å with a Z-score (measure of spatial significance of the fit relative to the alignment of random structures) of 6.7 and a gap size of 18 amino acids. The structure of Rad50, a DNA-binding protein where a structure analogous to that of a HisP monomer is formed from two domains, was also examined (15). An overlay of HisP and Rad50 is shown in Fig. 7B. The sequence identity between Rad50 and HisP is 20.2%, but the RMSD for alpha -carbon atoms is 2.7 Å, with a Z-score of 5.3 and a gap size of 29 amino acids. A sequence alignment of MalK, HisP, NBD1, NBD2, and Rad50 is shown in Fig. 8, which also shows the alignment of secondary structural elements for MalK, HisP, and Rad50. These results suggest that NBD1 and NBD2, with sequence identities to HisP of 18% and 17% and gap sizes of 32 and 17, respectively, may have similar three-dimensional structures to those of HisP, MalK, and Rad50. Figure 8 also indicates epitopes in NBD1 that bind NBD2 (blue), epitopes in NBD2 that bind NBD1 (red), and epitopes in either NBD that bind to the R-domain (green). Whereas the monomeric structures of these three proteins are very similar, the crystallographic dimers (Fig. 9) show no similarity. In addition to the crystallographic dimers for HisP, MalK, and Rad50, the comparison of NBD sequences from a large number of ABC transporters has led to the development of an alternative model for the HisP dimer, aHisP (19). In Fig. 9, A-D, the aqua monomer (NBD1) is always in the same orientation, and the arrow indicates the orientation of the helix in NBD2 that is projecting toward the viewer in Fig. 9A. Epitopes from one NBD of CFTR that bind to the other NBD are indicated with a darker color. Because they are found in the interfacial region of the crystal structures for MalK (9C), Rad50 (9D), and aHisP (9B), our data are more consistent with these models than with the HisP crystal structure (9A).


View larger version (52K):
[in this window]
[in a new window]
 
Fig. 7.   Alignment of NBD crystal structures. A: the crystal structure of the HisP monomer (17) is aligned to that for MalK (7). B: the HisP structure is aligned with one NBD of the Rad50 dimer (15). In A and B, HisP is shown in aqua and bound ATP in green. MalK and Rad50 are indicated in purple, with 2 tints used to indicate the 2 subunits that comprise the Rad50 NBD. Bound pyrophosphate (MalK) and ATP (Rad50) pyrophosphate are shown in red. The alignments were generated by minimizing the root mean square deviation between the alpha -carbons of the Walker A lysine and Walker B aspartate (amino acids 45 and 178 in HisP).



View larger version (42K):
[in this window]
[in a new window]
 
Fig. 8.   Sequence alignment of MalK, HisP, NBD1, NBD2, and Rad50. Sequence alignments are taken from published comparisons of MalK and HisP (7); HisP, NBD1, and NBD2 (17); and Rad50, NBD1, and NBD2 (15). Two large segments of Rad50 that have no counterparts in NBDs from ABC transporters are indicated below the alignment in lower case letters. Secondary structure assignments for MalK, HisP, and Rad50 are indicated by the colored amino acids with red indicating beta -sheet and aqua indicating alpha -helix. Walker A, B, and C motifs, as well as the Q-, D-, and H-loops, are highlighted in yellow. Epitopes in NBD1 that bind to NBD2 are highlighted in blue. Epitopes in NBD2 that bind to NBD1 are highlighted in red. Epitopes in either NBD that bind R-domain are highlighted in green.



View larger version (80K):
[in this window]
[in a new window]
 
Fig. 9.   Localization of peptides that bind to NBD1, NBD2, or R-domain in dimeric structures of HisP, MalK, and Rad50. Ribbon drawing for the dimeric crystal structures of HisP (17), MalK (7), and Rad50 (15) are shown in A, C, and D. E, G, and H show space-filling models of the same structures. B and F show the aHisP structure (19). In A-H, NBD1 (aqua) is shown in the same orientation. In B-D, the arrow indicates the orientation of the helix (X) in A, whose COOH-terminus is facing the viewer. In A-D, ATP (A, B, D), or pyrophosphate (C) is shown in yellow, epitopes in NBD1 that bind NBD2 are indicated in red, and epitopes in NBD2 that bind NBD1 are indicated in dark blue. In E-H, peptides in either NBD that bind to the R-domain are indicated in green.

These models can also be used to identify the location of peptides in NBD1 and NBD2 that bind to the R-domain. This is a more speculative analysis because the model proteins do not contain structures that are analogous to the R-domain. However, because CFTR is a member of the ABC transport superfamily, it is reasonable to propose that the basic alignment of NBDs and MSDs in CFTR is similar to that for other ABC transport superfamily members. If this is the case, then the R-domain should associate on the surface of the basic structure formed by the NBDs and MSDs. To test this hypothesis, we mapped epitopes in NBD1 and NBD2 that bind the R-domain onto the structures in Fig. 9, E-H. The HisP and MalK crystal structures (E and G) suggest that binding sites for the R-domain in NBD1 and NBD2 form two isolated patches. In contrast, the Rad50 and alternate HisP structures (F and H) suggest that R-domain binds to NBD1 and NBD2 along a long stripe. Although we do not regard this as conclusive evidence for any structure, we suggest that the data favor the Rad50 and alternate HisP structures. However, it is possible that peptides derived from regions of NBD1 and NBD2 that lie between the indicated binding sites may not be retained on our membranes.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Since the discovery of the gene for cystic fibrosis and the identification of transmembrane helices with hydropathy plots (35), there has been little success in efforts to further refine the structure of CFTR. At present, there is consensus on the domain boundaries (3). Yeast two-hybrid studies suggest the presence of interactions between intracellular domains and intracellular loops in the transmembrane domains (22), and binding of the NH2 and COOH termini to the intracellular domains has been proposed on the basis of functional studies (29, 32, 42). Whereas the NBDs of CFTR have been expressed and shown by various groups to bind or hydrolyze ATP (16, 23, 31) and the conformation of expressed R-domain has been shown to be altered by phosphorylation (8), there is presently no structural information for any of these domains. In a recent study, the CD spectrum of an R-domain construct (amino acids 708-831) that lacks the first 119 amino acids of our construct was analyzed and found to have little defined secondary structure (30). This construct was shown to restore kinase-dependent channel activity to a CFTR construct that lacked amino acids 708-831, but the presence of the NH2-terminus region of the R-domain in the CFTR construct may induce bound R-domain to assume a native structure. A similar effect could occur in solution. If this were the case, results should be more consistent with earlier studies of an expressed R-domain construct from amino acids 595-831, in which considerably more secondary structure was observed and in which PKA-dependent phosphorylation caused an appreciable change in the CD spectrum (8). It has been suggested that expressed R-domain is largely unstructured in solution. Whereas an unstructured R-domain might bind to the NBDs and specific peptides, the binding data in Figs. 5 and 6 and the phosphorylation data in Fig. 3 would appear to be more consistent with an R-domain with a defined tertiary structure. We therefore suggest that the absence of amino acids from the NH2 terminus of the R-domain peptide may in large part cause the reported absence of R-domain structure.

At present, the best structural information for CFTR comes from studies of other ABC transporters. Crystallographic structures have been reported for NBDs from two bacterial ABC transporters, the HisP subunit of the S. typhimurium histidine permease (17) and the MalK subunit of the T. litoralis maltose transporter (7). In addition, a crystal structure for Rad50 from Pyroccus furiosus has also been solved (15). The structures of two additional NBDs, MJ1267 and MJ0796, have recently been reported (21, 44). The monomeric structures are very similar to those of MalK and HisP, but they have not been considered in this analysis because the authors chose not to infer dimeric structures from their crystallographic data. In addition, the recent structure of the ABC transporter homolog MsbA was not considered because much of the NBD is unresolved in the crystal (4). Also, the authors' conclusion that the NBDs may not interact appears to be dependent on a series of interactions between extracellular loops in the transmembrane domains. Because these loops are considerably shorter in CFTR (see Fig. 6A), it is highly unlikely that CFTR could form a structure similar to that described from MsbA.

As described in RESULTS, these proteins show little sequence similarity but a great deal of structural homology. The regions with the most highly conserved sequences are those indicated in Fig. 8: the Walker A, B, and C regions and the Q-, D-, and H-loops. These regions have been linked to the binding or hydrolysis of ATP (7, 15, 17). With regard to domain-domain interactions between the NBDs, the crystallographic studies have generated vastly different dimeric structures. This is perhaps to be expected as the conditions for NBD crystallization are quite different from those found within the cell. One difference is that the MSDs are not present during crystallization, and, therefore, any effect that these domains have on interactions between the NBDs will not be reflected in the crystal structure. A better model for the structure of the NBD dimer in CFTR may be the soluble Rad50 dimer, formed in the presence of ATP, in which alterations in structure due to the absence of MSDs should not occur. In general, there are two different models for NBD-NBD interaction. The HisP dimer places the ATP binding sites on opposite sides of the dimer (17), whereas the Rad50 and MalK structures place the ATP binding sites in a cleft formed by the NBD-NBD interface (7, 15). For both of these models, ATP binding sites are likely to be composed of residues from both NBDs. Whereas this issue may be resolved only with the crystal structure of a complete ABC transporter, cysteine mutagenesis of amino acids in the ATP binding sites of MDR allows disulfide cross-links to be generated between the NBDs (25). This result is consistent only with an interfacial model. An ATP binding site at the interface of an NBD-NBD dimer may also explain why it has been difficult to observe ATP hydrolysis by NBD constructs.

In this study, we expressed, purified, and refolded tagged NBD1, NBD2, NBD1-R, and R-domain. These expressed CFTR domains had properties that were consistent with a native-like structure. All NBD-containing proteins bound ATP. The affinities for ATP binding to his-NBD1 and his-NBD2 were similar to values reported previously. However, it is somewhat surprising that the affinity of NBD2 is greater than that of NBD1, because previous studies have suggested the opposite (23, 31). The only explanation we can offer is that, in the previous studies, NBD1 and NBD2 constructs from amino acids 433-589 and 1208-1399 were fused to the maltose binding protein, whereas our NBD1 and NBD2 constructs were His-tagged and spanned amino acids 373-589 and 1151-1476 (23, 31). Recent studies have indicated that NBD1 may extend as far as amino acid 640 (3). However, the observation that, when fused to the maltose-binding protein, an NBD1 construct that terminates at amino acid 589 can hydrolyze ATP (26) suggests that amino acids after 589 are not essential for the formation of a native-like structure. In vitro phosphorylation of NBD1-R by PKA occurred at the same sites as in situ phosphorylation of full-length CFTR. In addition, the sites of R-domain phosphorylation differed from those observed when immunoprecipitated CFTR was in vitro phosphorylated with PKA. Direct associations between the expressed domains were observed with overlay assays and by tryptophan fluorescence quenching. The dose dependence of tryptophan fluorescence quenching allowed binding to be quantified. However, since we have no data regarding homodimerization, caution must be used with regard to quantification of the association between NBD1-R and NBD2. Recently, it has been reported that dimers of NBDs are not formed in solution (21). This has been used as an argument in favor of the possibility that crystallographic NBD dimers may not reflect the structure within native ABC transporters. Although we are in agreement with this conclusion, our fluorescence quenching data indicate that NBDs can associate in solution. Perhaps the presence of the R-domain stabilizes the interaction, because an NBD1-R/NBD2 complex has been observed previously (26). We also suspect that conditions designed to precipitate NBDs may disfavor solution dimer formation, whereas ours, designed to maintain NBDs in solution, favored dimer formation. Differences in the size of the expressed domains may also affect domain-domain interactions. Lastly, we have shown that an association between NBD1-R and NBD2 alters the effects of ATP binding on tryptophan fluorescence.

The principle finding of our study is that specific epitopes in a peptide library spanning all cytoplasmic regions of CFTR bind to the expressed domains of CFTR. Epitopes from one domain that bind to another domain are likely to define sites of interaction between the two domains. Because the epitopes in one NBD that bind to the other NBD are at the interface in models that place the ATP binding sites at the NBD-NBD interface, our data are most consistent with the crystal structures of MalK, Rad50, and the alternative structure for HisP. Our data are in agreement with crystallographic (7, 15) and cross-linking (25) studies of other ABC superfamily members. However, because there are complications with each approach, the confluence of data with three different techniques and with three different ABC superfamily members is comforting.

In the present study, care has been taken to exclude false positive results. Our ad hoc procedures for identifying false positives are described in RESULTS, but an additional test for specific binding made use of the fact that each six-amino acid sequence is expressed on three different peptides (see Fig. 6A). As a consequence, a series of sequential peptides that bound a domain was considered to be a stronger indication of a specific interaction than binding by a single peptide. However, since peptides may form structures that obscure the relevant epitope, strong binding by an isolated peptide cannot be ignored. Binding to sequential peptides also defines the site of interaction more precisely. In addition, the peptides that bound NBD1 and NBD2 were not the same. These observations strongly suggest that, in general, peptide binding to our expressed domains was specific.

In addition to interactions between NBD1 and NBD2, interactions between R-domain and both NBDs were observed. Previous functional studies of CFTR channel activity have suggested that the activities of the NBDs and R-domain are linked (18). In an NBD1-R-domain construct, R-domain phosphorylation inhibits nucleotide binding, and the presence of R-domain alters the kinetics of ATP hydrolysis by NBD1 (27, 43). Our results provide direct physical evidence for these interactions. The analysis of peptide binding to the R-domain in terms of NBD crystal structure is more speculative than that for NBD-NBD interaction because the model proteins do not have an R-domain. However, because we have based our analysis on the assumption that arrangement of NBDs and MSDs in ABC superfamily members are the same, regions of NBD1 and NBD2 that associate with R-domain should form a contiguous surface. Although none of the structures in Fig. 9 are entirely consistent with our assumption, the peptides that interact with the R-domain are most contiguous in the Rad50 and aHisP structures. The MalK and HisP crystals show binding of the R-domain at two distant locations. On the basis of these observations, we conclude that the Rad50 structure and the alternative aHisP (20) structures represent the most likely structures for the NBDs in CFTR.


    ACKNOWLEDGEMENTS

The technical assistance of Mu-Young Kim is acknowledged.


    FOOTNOTES

This work was supported by grants from the Cystic Fibrosis Foundation, Cystic Fibrosis Research, and the Nemours Foundation.

Address for reprint requests and other correspondence: W. W. Reenstra, Institute for Human Gene Therapy, Univ. of Pennsylvania, Philadelphia, PA 19104 (E-mail: Reenstra{at}mail.med.upenn.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.

First published January 2, 2002;10.1152/ajpcell.00337.2001

Received 20 July 2001; accepted in final form 27 November 2001.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Anderson, MP, Berger HA, Rich DR, Gregory RJ, Smith AE, and Welsh MJ. Nucleoside triphosphates are required to open the CFTR chloride channel. Cell 67: 775-784, 1991[ISI][Medline].

2.   Boyle, WJ, van der Geer P, and Hunter T. Phosphopeptide mapping and phosphoamino acid analysis by two-dimensional separation on thin-layer plates. Methods Enzymol 210: 110-148, 1991.

3.   Chan, KW, Csanady L, Seto-Young D, Nairn AC, and Gadsby DC. Severed molecules functionally define the boundaries of the cystic fibrosis transmembrane conductance regulator's NH2-terminal nucleotide binding domain. J Gen Physiol 116: 163-180, 2000[Abstract/Free Full Text].

4.   Chang, G, and Roth CB. Structure of MsbA from E. coli: a homolog of the multidrug resistance ATP binding cassette (ABC) transporters. Science 293: 1793-1800, 2001[Abstract/Free Full Text].

5.   Cheng, SH, Rich DP, Marshall J, Gregory RJ, Welsh MJ, and Smith AE. Phosphorylation of the R domain by cAMP-dependent protein kinase regulates the CFTR chloride channel. Cell 66: 1027-1036, 1991[ISI][Medline].

6.   Chen, JH, Chang XB, Aleksandrov AA, Hammerk MM, Shanmugam K, and Riordan JR. Biochemical and electrophysiological evaluation of CFTR quaternary structure (Abstract). Pediatr Pulmonol Suppl 20: 174, 2000.

7.   Diederichs, K, Diez J, Greller G, Muller C, Breed J, Schnell C, Vonrhein C, Boos W, and Welte W. Crystal structure of MalK, the ATPase subunit of the trehalose/maltose ABC transporter of the archaeon Thermococcus litoralis. EMBO J 19: 5951-5961, 2000[Abstract/Free Full Text].

8.   Dulhanty, AM, and Riordan JR. Phosphorylation by cAMP-dependent protein kinase causes a conformational change in the R-domain of the cystic fibrosis conductance transmembrane regulator. Biochemistry 33: 4072-4079, 1994[ISI][Medline].

9.   Eskandari, S, Wright EM, Kreman M, Starace DM, and Zampighi GA. Structural analysis of cloned plasma membrane proteins by freeze-fracture electron microscopy. Proc Natl Sci USA 95: 11235-11240, 1998[Abstract/Free Full Text].

10.   Frangioni, JV, and Neel BG. Solubilization and purification of enzymatically active glutathione S-transferase (pGEX) fusion proteins. Anal Biochem 210: 179-187, 1993[ISI][Medline].

11.   Gadsby, DC, and Nairn AC. Control of CFTR channel gating by phosphorylation and nucleotide hydrolysis. Physiol Rev 79: S77-S107, 1999[Medline].

12.   Hall, RA, Ostedgaard LS, Premont RT, Blitzer JT, Rahman N, Welch MJ, and Lefkowitz RJ. A C-terminal motif found in the beta2-adrenergic receptor, P2Y1 receptor, and cystic fibrosis transmembrane conductance regulator determines binding to the Na+/H+ exchanger regulatory factor family of PDZ proteins. Proc Natl Acad Sci USA 95: 8496-8501, 1998[Abstract/Free Full Text].

13.   Hallows, KR, Raghuram V, Kemp BE, Witters LA, and Foskett JK. Inhibition of cystic fibrosis transmembrane conductance regulator by novel interaction with the metabolic sensor AMP-activated protein kinase. J Clin Invest 105: 1711-1721, 2000[Abstract/Free Full Text].

14.   Higgins, CF. ABC transporters: from microorganisms to man. Annu Rev Cell Biol 8: 67-113, 1992[ISI].

15.   Hopfner, KP, Karcher A, Shin DS, Craig L, Arthur LM, Carney JP, and Tainer JA. Structural biology of Rad50 ATPase: ATP-driven conformational control in DNA double-strand break repair and the ABC-ATPase superfamily. Cell 101: 789-800, 2000[ISI][Medline].

16.   Howell, LD, Borchardt R, and Cohn JA. ATP hydrolysis by a CFTR domain: pharmacology and effects of G551D mutation. Biochem Biophys Res Commun 271: 518-525, 2000[ISI][Medline].

17.   Huang, LW, Wang IX, Nikaido K, Liu PQ, Ames GFL, and Kim SH. Crystal structure of the ATP-binding subunit of an ABC transporter. Nature 396: 703-707, 1998[ISI][Medline].

18.   Hwang, TC, Wang F, Yang CH, and Reenstra WW. Potentiation of Delta F508 channel function by genistein binding to CFTR. Am J Physiol Cell Physiol 273: C988-C998, 1997[Abstract/Free Full Text].

19.   Jones, PM, and George AM. Subunit interactions in ABC transporters: towards a functional architecture. FEMS Microbiol Lett 179: 187-200, 1999[ISI][Medline].

20.   Joseph, G, and Pick E. "Peptide walking" is a novel method for mapping functional domains in proteins. Its application to the Rac1-dependent activation of NADPH oxidase. J Biol Chem 270: 29079-29082, 1995[Abstract/Free Full Text].

21.   Karpowich, N, Martsinkevich O, L, Millen Yuan YR, Dei PL, MacVey K, Thomas P, and Hunt JF. Crystal structures of the MJ1267 ATP-binding cassette reveal an induced-fit effect at the ATPase active site of an ABC transporter. Structure 9: 571-586, 2001[ISI][Medline].

22.   Kiser, GL, Chang XB, and Riordan JR. Two-hybrid analysis of CFTR domain interactions (Abstract). Pediatr Pulmonol Suppl 13: 213, 1996.

23.   Ko, YH, and Pedersen PL. The first nucleotide binding fold of the cystic fibrosis transmembrane conductance regulator can function as an active ATPase. J Biol Chem 270: 22093-22096, 1995[Abstract/Free Full Text].

24.   Kyte, J, and Doolittle RF. A simple method for displaying the hydropathic character of a protein. J Mol Biol 157: 105-132, 1982[ISI][Medline].

25.   Loo, TW, and Clarke DM. Covalent modification of human P-glycoprotein mutants containing a single cysteine in either nucleotide-binding fold abolishes drug-stimulated ATPase activity. J Biol Chem 270: 22957-22961, 1995[Abstract/Free Full Text].

26.   Lu, NT, and Pedersen PL. Cystic fibrosis transmembrane conductance regulator: the purified NBF1+R protein interacts with the purified NBF2 domain to form a stable NBF+R/NBF2 complex while inducing a conformational change transmitted to the C-terminal region. Arch Biochem Biophys 375: 7-20, 2000[ISI][Medline].

27.   Ma, J, Zhao J, Drumm ML, Xie J, and Davis PB. Function of the R-domain in the cystic fibrosis transmembrane conductance regulator chloride channel. J Biol Chem 272: 28133-28141, 1997[Abstract/Free Full Text].

28.   Marshall, J, Fang S, Ostedgaard LS, O'Riordan CR, Ferrara D, Amara JF, Hoppe H, Scheule RK, Welsh MJ, and Smith AE. Stoichiometry of recombinant cystic fibrosis transmembrane conductance regulator in epithelial cells and its functional reconstitution into cells in vitro J. Biol Chem 269: 2987-2995, 1994[Abstract/Free Full Text].

29.   Naren, AP, Quick MW, Collawn JF, Nelson DJ, and Kirk KL. Syntaxin 1A inhibits CFTR chloride channels by means of domain-specific protein-protein interactions. Proc Natl Acad Sci USA 95: 10972-10977, 1998[Abstract/Free Full Text].

30.   Ostedgaard, LS, Baldursson O, Vermeer DW, Welsh MJ, and Robertson AD. A functional R-domain from cystic fibrosis transmembrane conductance regulator is predominantly unstructured in solution. Proc Natl Acad Sci USA 97: 5657-5662, 2000[Abstract/Free Full Text].

31.   Randak, C, Neth P, Auerswald EA, Eckerskorn C, Assfalg-Machleidt I, and Machleidt W. A recombinant polypeptide model of the second nucleotide-binding fold of the cystic fibrosis transmembrane conductance regulator functions as an active ATPase, GTPase, and adenylate kinase. FEBS Lett 410: 180-186, 1997[ISI][Medline].

32.   Raghuram, V, Mak DD, and Foskett JK. Regulation of cystic fibrosis transmembrane conductance regulator single-channel gating by bivalent PDZ-domain-mediated interaction. Proc Natl Acad Sci USA 98: 1300-1305, 2001[Abstract/Free Full Text].

33.   Reenstra, WW, Yurko-Mauro K, Dam A, Raman S, and Shorten S. CFTR chloride channel activation by genistein: the role of serine/threonine protein phosphatases. Am J Physiol Cell Physiol 271: C650-C657, 1996[Abstract/Free Full Text].

34.   Rich, DP, Berger HA, Cheng SH, Travis SM, Saxena M, Smith AE, and Welsh MJ. Regulation of the cystic fibrosis transmembrane conductance regulator Cl- channel by negative charge in the R-domain. J Biol Chem 268: 20259-20267, 1993[Abstract/Free Full Text].

35.   Riordan, JR, Rommens JM, Kerem B, Alon N, Rozmahel R, Grzelczak Z, Zielenski J, Lok S, Plavsic N, Chou JL, Drumm ML, Iannuzzi MC, Collins FS, and Tsui LC. Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 245: 1066-1073, 1989[ISI][Medline].

36.   Seibert, FS, Chang XB, Aleksandrov AA, Clarke DM, Hanrahan JW, and Riordan JR. Influence of phosphorylation by protein kinase A on CFTR at the cell surface and endoplasmic reticulum. Biochem Biophys Acta 1461: 275-283, 1999[ISI][Medline].

37.   Sheppard, D, and Welsh MJ. Structure and function of the CFTR chloride channel. Physiol Rev 79: S23-S45, 1999[Medline].

38.   Shindyalov, IN, and Bourne PE. Protein structure alignment by incremental combinatorial extension (CE) of the optimal path. Protein Eng 11: 739-747, 1998[Abstract].

39.   Short, DB, Trotter KW, Reczek D, Kreda SM, Bretscher A, Boucher RC, Stutts MJ, and Milgram SL. An apical PDZ protein anchors the cystic fibrosis transmembrane conductance regulator to the cytoskeleton. J Biol Chem 273: 19797-19801, 1998[Abstract/Free Full Text].

40.   Smith, PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, Fujimoto EK, Goeke NM, Olson BJ, and Klenk DC, DC Measurement of protein using bicinchoninic acid. Anal Biochem 150: 76-85, 1985[ISI][Medline].

41.   Valerio, RM, Benstead M, Bray AM, Campbell RA, and Maeji NJ. Synthesis of peptide analogs using the multipin peptide synthesis method. Anal Biochem 197: 168-177, 1991[ISI][Medline].

42.   Wang, S, Yue H, Derin RB, Guggino WB, and Li M. Accessory protein facilitated CFTR-CFTR interaction, a molecular mechanism to potentiate the chloride channel activity. Cell 103: 169-179, 2000[ISI][Medline].

43.   Winter, MC, and Welsh MJ. Stimulation of CFTR activity by its phosphorylated R-domain. Nature 389: 294-296, 1997[ISI][Medline].

44.   Yuan, YR, Blecker S, Martsinkevich O, Millen L, Thomas PL, and Hunt JF. The crystal structure of the MJ0796 ATP-binding cassette: implications for the structural consequences of ATP hydrolysis in the active site of an ABC-transporter. J Biol Chem 276: 32313-32321, 2001[Abstract/Free Full Text].

45.   Yurko-Mauro, KA, and Reenstra WW. Prostaglandin F2alpha stimulates CFTR activity by PKA and PKC-dependent phosphorylation. Am J Physiol Cell Physiol 275: C653-C660, 1998[Abstract].

46.   Zeltwanger, S, Wang F, Wang GT, Gillis KD, and Hwang TC. Gating of cystic fibrosis transmembrane conductance regulator chloride channels by adenosine triphosphate hydrolysis. Quantitative analysis of a cyclic gating scheme. J Gen Physiol 113: 541-554, 1999[Abstract/Free Full Text].

47.   Zerhusen, B, Zhao J, Xie J, Davis PB, and Ma J. A single conductance pore for chloride ions formed by two cystic fibrosis transmembrane conductance regulator molecules. J Biol Chem 274: 7627-7630, 1999[Abstract/Free Full Text].


Am J Physiol Cell Physiol 282(5):C1170-C1180
0363-6143/02 $5.00 Copyright © 2002 the American Physiological Society