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Review

On the structural organization of the intracellular domains of CFTR夽 Oscar Moran ∗ Instituto di Biofisica, CNR, Via de Marini, 6, 16149 Genova, Italy

a r t i c l e

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Article history: Received 15 October 2013 Received in revised form 17 January 2014 Accepted 21 January 2014 Available online xxx Keywords: CFTR NBD Regulatory domain Cystic fibrosis Structure

a b s t r a c t The cystic fibrosis transmembrane conductance regulator (CFTR) is a multidomain membrane protein forming an anion selective channel. Mutations in the gene encoding CFTR cause cystic fibrosis (CF). The intracellular side of CFTR constitutes about 80% of the total mass of the protein. This region includes domains involved in ATP-dependent gating and regulatory protein kinase-A phosphorylation sites. The high-resolution molecular structure of CFTR has not yet been solved. However, a range of lower resolution structural data, as well as functional biochemical and electrophysiological data, are now available. This information has enabled the proposition of a working model for the structural architecture of the intracellular domains of the CFTR protein. This article is part of a Directed Issue entitled: Cystic Fibrosis: From o-mics to cell biology, physiology, and therapeutic advances. © 2014 Elsevier Ltd. All rights reserved.

Contents 1. 2. 3. 4.

5.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The structure of the nucleotide binding domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The structure of the regulatory domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Intracellular inter-domain organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. NBD1–NBD2 interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. NBDs–ICLs interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. NBDs–RD interactions and phosphorylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction Cystic fibrosis transmembrane conductance regulator (CFTR; ATP-binding cassette (ABC) sub-family C, member 7, Uniprot P13569) is an integral membrane protein expressed mainly in vertebrate epithelia. In humans, CFTR mutations are associated with cystic fibrosis (CF), the most common lethal genetic disease (for reviews see Welsh and Ramsey, 1998; Quinton, 1999; O’Sullivan and Freedman, 2009; Goss and Ratjen, 2013). CFTR functions as an anionic channel that regulates anion, mainly chloride and bicarbonate, transport in epithelia (Akabas, 2000; Frizzell and Hanrahan, 2012).

夽 This article is part of a Directed Issue entitled: Cystic Fibrosis: From o-mics to cell biology, physiology, and therapeutic advances. ∗ Tel.: +39 0106475558. E-mail address: [email protected]

The general topology of CFTR, illustrated in Fig. 1, consists of two membrane-spanning domains (MSD), each with six transmembrane helices, two nucleotide-binding domains (NBD) and a regulatory domain (RD).1 The intracellular moiety of CFTR, NBD1, NBD2, RD, N-terminus, C-terminus, and four intracellular linkers of the transmembrane helices (ICL1–ICL4) represents 80% of the total amino acid content of the protein (1164 out of 1480 residues). These domains are responsible for the ATP-dependent gating of CFTR (NBD1 and NBD2) and protein kinase-A regulation of the channel (RD). It is noteworthy that about one third of the CFTR-mutations correlated with CF are located in the intracellular side of the protein (see www.genet.sickkids.on.ca). Indeed, 9 out of the 10 more frequent mutations, including the worldwide most common

1 Boundaries of the human CFTR regions and domains are taken as proposed in the Uniprot database (http://www.uniprot.org), entry P13569.

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Fig. 1. The topology of CFTR. The scheme is depicted in a plausible position respect to the plasma membrane. MSD1 and MSD2 are the membrane spanning domains; NBD1 and NBD2 are the nucleotide binding domains; RD is the regulatory domain; ICL1 to ICL4 are the intracellular loops 1–4.

mutation, deletion of the residue phenylalanine 508 (F508del), occur in the intracellular side of the protein (Bobadilla et al., 2002). Defining the molecular structure of CFTR at high resolution by means of X-ray crystallography is a highly complicate task for a number of reasons. CFTR is a high molecular mass (∼168 kDa) membrane protein, which poses considerable problems for purification and solubilization. As a huge amphoteric protein, that includes an intrinsically disordered region, it is not a good candidate for forming crystals. On the other hand, the size of CFTR limits the possibility of solving its atomic structure by NMR methods. Therefore, the molecular structure of the whole protein has been successfully studied by low-resolution electron microscopy methods (Awayn et al., 2005; Mio et al., 2008; Zhang et al., 2009, 2011; Rosenberg et al., 2004, 2011; Ford et al., 2011), while high resolution methods have been limited only to NBD1 and NBD2. 2. The structure of the nucleotide binding domains The nucleotide-binding domains of ABC proteins are known to bind and hydrolyse ATP, thereby coupling transport to ATP hydrolysis in a large number of biological processes. In CFTR, NBD1 is defined as the sequence between residue 423 and 646, and NBD2 between residue 1210 and residue 1443. The primary structure of the NBDs is duplicated in several ABC-protein subfamilies (Dassa and Bouige, 2001; Mendoza and Thomas, 2007). Indeed, both nucleotide-binding domains, NBD1 and NBD2, are the most conserved portions of CFTR with almost full protein sequence identity for most primates, remarkably high identity for other mammals (>90%), and slightly lower identity for most other vertebrates (>80%). Conversely, in CFTR NBD1 and NBD2 share lower sequence similarity (∼30%). Residue conservation of NBD sequences between species has allowed the development of homology-based molecular models of the NBDs (Bianchet et al., 1997; Callebaut et al., 2004; Moran et al., 2005). Several successful protocols have been applied to produce recombinant NBDs. Both NBD1 and NBD2 have been expressed in Escherichia coli, and purified either from the soluble fraction (Logan et al., 1994; Randak et al., 1995, 1997; Howell et al., 2000) or from the inclusion bodies (Hartman et al., 1992; Ko and Pedersen, 1995; Clancy et al., 1997; Neville et al., 1998; Lu and Pedersen, 2000; Wang et al., 2002; Galeno et al., 2011; Galfrè et al., 2012). Recombinant NBDs have also been produced in insect cells (Kidd et al., 2004). Recombinant NBD proteins obtained with these methods bind and hydrolyse ATP, have circular dichroism and UV spectra characteristic of folded NBDs and demonstrate domain–domain interaction patterns. However, as the native preparation of NBD1

and NBD2 tend to precipitate at relatively low concentration (>2.5 mg/ml; Galeno et al., 2011; Galfrè et al., 2012), to obtain protein concentrations compatible with the crystallization conditions, three to seven revertant mutations (F409L, F429S, F433L, G550E, R553Q, R555K, H667R, Roxo-Rosa et al., 2006; F429S, F494N, Q637R, Pissarra et al., 2008) have been introduced into the NBD1. These mutations, that increase NBD1 solubility, have allowed diffracting crystals to be obtained that solve the atomic structure of this domain (Lewis et al., 2004, 2005, 2010; Thibodeau et al., 2005; Atwell et al., 2010; Mendoza et al., 2012). Also the structure of NBD2 has been solved by X-ray crystallography (PDB entry 3GD7). The local architecture of the NBDs of CFTR (Fig. 2) is consistent with the known structures of sequence-related nucleotide-binding proteins (Jones and George, 1999; Callebaut et al., 2004). The nucleotide-binding domains are composed of two distinct subdomains, the catalytic core sub-domain (grey in Fig. 2A), and a smaller ␣-helical sub-domain (light yellow in Fig. 2A). The larger catalytic core sub-domain consists of three groups of ␤-sheets and six ␣-helices (see Fig. 2B and C). It contains two short, highly conserved consensus sequences, the Walker A and B motifs (Walker et al., 1982). The phosphate-binding (P-loop) or Walker A motif (GXXGXGKS/T) is a glycine-rich loop followed by an uncapped Khelix. It forms a flexible loop between a ␤-strand and an ␣-helix. Conserved lysine and serine residues within this structure interact with the phosphate groups and the magnesium ion of the bound Mg2+ -nucleotide complex. The Walker B motif is h-h-h-h-D, where ‘h’ is a hydrophobic residue. The conserved aspartate residue coordinates the catalytic Mg2+ ion. The helical sub-domain consists of four ␣-helices, that contain the ABC signature motif, also known as the LSGGQ motif, linker peptide or C motif. The LSGGQ motif is present in the NBD1, but altered to LSHGH in the NBD2. The NBDs also have a glutamine residue residing in a flexible loop called the Q loop, lid or ␥-phosphate switch, that connects the ␣-helical sub-domain and the catalytic core sub-domain. The Q-loop interacts with the ATP␥ phosphate through a water bond. The interaction of the adenyl moiety of ATP with an aromatic residue, phenylalanine 430 in NBD1 and tyrosine 1219 in NBD2, is a key determinant of nucleotide binding. The H-loop region contains a highly conserved histidine residue, that is postulated to polarize the attached water molecule for hydrolysis. The structure of NBD1 has an important variation with respect to other members of the ABC-transporter super-family (Lewis et al., 2004, 2005, 2010). There is a 35-residue (403–437) region that contains serine residues in phosphorylation motifs, named the regulatory insertion (RI). The RI constitutes a loop on the domain surface. This region is disordered in NBD1 crystals, and is not resolved in the electron density map. Phosphorylation of serine 422 orders the structure of the RI (Lewis et al., 2004). It has been proposed that phosphorylation of the RI may have a physiological role. However, deletion of a central segment (415–432) of the RI preserves CFTR function (Chan et al., 2000; Aleksandrov et al., 2010). Interestingly, deletion of the complete RI sequence promoted the maturation of F508 CFTR, perhaps modifying the interactions of NBD1 with the rest of the protein (Aleksandrov et al., 2010). The regulatory extension (RE), defined as residues 638–670, has also been described as a part of NBD1 (Lewis et al., 2004, 2005; Thibodeau et al., 2005; Mendoza et al., 2012). This region contains two serine residues that form part of phosphorylation motifs (660 and 670). After treatment with protein kinase-A (PKA), these two serines are phosphorylated. It is, however, disputable whether to include the RE as part of NBD1 or the RD. Indeed, the boundaries of NBD1 are still a controversial matter: the C-terminus limit has been defined as residue 586 (Riordan et al., 1989), residue 684 (Bianchet et al., 1997), residue 633 (Chan et al., 2000) or residue 646

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Fig. 2. Structure of the nucleotide binding domains. (A) Solvent accessible surface of the atomic structure of the human NBD1 (pdb 2PZE) and NBD2 (pdb 3GD7). The catalytic core is grey and the ␣-helical sub-domain is light-yellow. (B) The cartoon representation is rotated by 90◦ as indicated. Colours in the cartoons correspond to the conserved motifs: the LSGGQ-motif is yellow, the Walker-A is blue, the Walker-B is green, the H-loop is magenta, the Q-loop is red, and the switch-II motif is orange. (C) Scheme of the internal topology of the NBD’s motifs. The ␣-helices are represented as cylinders, and the ␤-strands as arrows. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of the article.)

(http://www.uniprot.org, entry P13569). Indeed, several authors consider that the RE forms part of the RD (Csanády et al., 2005; Baker et al., 2007; Kanelis et al., 2010; Marasini et al., 2012, 2013). Crystallographic solution of the structure of NBD1 has shown that mutation F508del does not introduce a significant difference to the domain fold, just a small difference in the surface near to the position of F508 (Lewis et al., 2005; Thibodeau et al., 2005). Molecular dynamics applied to the crystallographic data suggested that F508del–NBD1 has increased flexibility (Wieczorek and Zielenkiewicz, 2008), with local changes in the solvent accessibility around the mutation location (Bisignano and Moran, 2010). Local difference was also supported experimentally using HDXMS analysis (Lewis et al., 2010). The thermodynamic stability of the solubilized F508del–NBD1 is reduced (Mendoza et al., 2012; Thibodeau et al., 2005; Protasevich et al., 2010; Wang et al., 2010). This instability of the isolated NBD1 has been correlated with the inability of mutant CFTR to be targeted to the membrane. However, the precise mechanisms for the failed maturation of F508del-CFTR are not completely understood. 3. The structure of the regulatory domain The regulatory domain (RD) of CFTR is about 200 residues long, with 18 potentially phosphorylation sites (12 serines and 8 threonines). The sequence is highly conserved in mammals (>85% identity), but less so in other vertebrates (from 60% to 35% identity). There is no other related protein for which the atomic structure has been solved; thus homology modelling of CFTR’s RD is not feasible. Attempts to predict the RD structure by ab initio coarsegrain modelling (Hegedus et al., 2008) or by hydrophobic cluster analysis (Mornon et al., 2008, 2009) have yielded different, divergent molecular models. Cryo-electron microscopy observations of

anti-regulatory domain anti-body complex have shown that the RD occupies a position that extends from the bottom of the cytosolic regions up to the transmembrane domains (Mio et al., 2008; Zhang et al., 2009, 2011). The structure of the RD is predicted to be mostly disordered (Ostedgaard et al., 2001; Marasini et al., 2013). The intrinsic disorder of the RD is reflected in measurements of the dimensions of the isolated RD by size exclusion chromatography (SEC), small-angle X-ray scattering (SAXS) and light scattering (Marasini et al., 2013), that show an excess of the gyration radius estimation consistent with a molten protein (Uversky et al., 2000). Nevertheless, despite being intrinsically disordered, the isolated RD protein in solution retains a degree of secondary structure organization, as observed by circular dichroism (Dulhanty and Riordan, 1994; Marasini et al., 2012, 2013) or NMR (Baker et al., 2007; Kanelis et al., 2010). The RD plays a cardinal role in the regulation of CFTR by second messengers. Past experiments demonstrated that CFTR missing amino acids 708–835 (R-CFTR) led to PKA-independent activity, indicating that this region of the R-domain encodes residues necessary for PKA-dependent activation (Rich et al., 1991, 1993; Ma et al., 1997; Chang et al., 1993). In a fully phosphorylated protein, eight phosphoserines (residue positions 660, 700, 712, 737, 753, 768, 795 and 813) have been detected by mass spectrometry (Neville et al., 1997; Townsend et al., 1996) and NMR (Baker et al., 2007), and partial phosphorylation (approximately 60%) of serine at position 670 (Baker et al., 2007). There are several lines of evidence that suggest not all sites contribute equally to channel regulation. A large reduction in channel activity is caused by mutagenesis of only four sites (serines 660, 737, 795, and 813) (Wilkinson et al., 1997). When these sites as well as all of the other dibasic consensus sites were substituted (serines 686, 700, 712, 768, and threonine 788) activity was further reduced (Wilkinson et al., 1997), but additional

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monobasic sites were still able to support a low level of activity (Dulhanty and Riordan, 1994). Two of the dibasic sites at serines 737 and 768 have been described as distinctly different from the others, playing inhibitory rather than stimulatory roles when phosphorylated (Vais et al., 2004; Csanády et al., 2005). In the isolated RD, phosphorylation leads to conformational changes of the protein. The secondary structure composition is significantly altered when the protein is phosphorylated (Dulhanty and Riordan, 1994; Marasini et al., 2012, 2013; Baker et al., 2007; Kanelis et al., 2010). The conformational change of the RD following phosphorylation is also detectable by intrinsic fluorescence spectroscopy assays (Marasini et al., 2013). This change is accompanied by reduction of the Strokes radius of the RD (from 3.6 nm to 3.4 nm) measured by SEC (Marasini et al., 2013). Thermodynamic analysis of denaturation curves of the native and phosphorylated RD showed that phosphorylation of the protein induces a state of lower stability (Marasini et al., 2012). SAXS measurements confirmed the condensation of the RD upon phosphorylation. The Guinier analysis of SAXS data yielded a gyration radius, Rg , of 3.25 nm for the native RD, and 2.92 nm for the phosphorylated RD (Marasini et al., 2013). The maximum size of the molecule, Dmax , estimated from the pair distance distribution function (PDDF) gave a value of 11.4 nm for the native RD, and 10.2 nm for the phosphorylated polypeptide (Marasini et al., 2013). The shape of the PDDF obtained from the SAXS data of RD in the two states are slightly different, with maxima that are smaller than Dmax /2 (Fig. 3A), which correspond to elongated molecules. Thus, these differences in the size may be due to a change in the shape of the molecule, as suggested by comparing the PDDF presented in Fig. 3A. As an intrinsically disordered protein, the RD is characterized by lack of stable tertiary structure (Uversky et al., 2000; Dunker et al., 2002). Obviously, intrinsically disordered proteins cannot be crystallized. An improved quantitative characterization of flexible and intrinsically disordered proteins in solution can be obtained from SAXS data, assuming that the experimental data can be represented by an average of a (generally unknown) number of conformers (Bernadó and Svergun, 2012). Thus, a pool containing a large number of possible conformations is randomly generated to cover the configurational space, and an ensemble optimization method (EOM) is used to select appropriate subsets of configurations fitting the experimental data. Applying the EOM to the experimental SAXS data of the proteins in solution, ensembles of the native and phosphorylated RD (Fig. 3B and C) were obtained (Marasini et al., 2013). Ensembles were characterized by an anisometry larger than 1, which is consistent with the elongated prolate shape of the molecules. The EOM analysis also indicated differences between the ensembles for the native and phosphorylated RD. Indeed, the phosphorylated RD has a smaller average Rg and Dmax than the native protein (Fig. 3D and E). Interestingly, the prediction of the structure of the native and phosphorylated RD by coarse-grain molecular dynamics simulations (Hegedus et al., 2008) resulted in an ensemble with a smaller gyration radius for the native (1.82 nm in simulation vs. 3.36 nm from experimental data), and predicted an increase in size when the protein is phosphorylated (1.97 nm), while a compaction of the molecules is observed from experimental data (2.97 nm; Marasini et al., 2013). Some disordered regions of proteins can form ordered structures when they bind to targets (Marsh et al., 2010). In fact, when the regulatory extension, RE (residues 638–676), is crystallized together with NBD1, it has a strong tendency to form an amphipathic helix that docks at a preferential site on the surface of NBD1 (Lewis et al., 2004). Therefore, the possibility cannot be ruled out that when the RD forms part of the whole CFTR protein, the interactions with other protein domains may restrict the degree of disorder observed in the isolated protein in solution. It is important to note that the conformational changes observed in the isolated

Fig. 3. Structural features of the CFTR-regulatory domain. (A) The pair distance distribution function of the native (black) and phosphorylated (red) RD calculated from the SAXS data obtained from the two protein preparations in solution. Representation of 16 conformations of the native RD (B) and 19 conformations of the phosphorylated RD (C) selected by the EOM analysis, that best-fit the SAXS data, yielding the two subsets of distribution of the Rg (D) and Dmax (E). The distributions of the random pool are shown in black, those of the selected ensemble of native RD are in light blue and those of phosphorylated RD in red. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of the article.)

domains may be related to different interactions of the RD with the NBDs in the activated and inactivated CFTR. It is conceivable that the conformational variability of the RD offers it the advantage of adaptive binding to different targets within the CFTR protein. 4. Intracellular inter-domain organization A general scheme of CFTR function can be summarized as follows: activation of CFTR occurs upon PKA-dependent phosphorylation of the RD, and gating of the CFTR channel comes after binding of ATP to the NBDs. This sequence of events implies a series of conformational changes that, in turn condition the successive step. Phosphorylation of the RD produces a conformational change of the domain (see Section 3) that favours the binding of ATP to

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Fig. 4. Dimeric conformation of NBD1 and NBD2. (A) The dimer formed by NBD1 (light blue) and NBD2 (pink) is represented with ATP molecules in the binding site. The Walker A (blue) of NBD2 and ABC signature (yellow) of NBD1 forms the NBD1 composite binding site, and the Walker A of NBD1 and ABC signature of NBD2 forms the NBD2 composite binding site. The low-resolution SAXS-reconstruction of the NBD1–NBD2 dimer in the absence (B) and presence (C) of ATP. Molecular homology-models of the NBD1–NBD2 dimer, in the closed (B) and open (C) channel conformations (Mornon et al., 2009) are docked to the SAXS-based models (Galeno et al., 2011). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of the article.)

the NBDs (Winter and Welsh, 1997; Mathews et al., 1998; Csanády et al., 2000), which is likely to induce the formation of an ATPdriven tight NBD1–NBD2 heterodimer (Vergani et al., 2003, 2005; Mense et al., 2006). Analogous to what occurs to other ABC proteins (Dawson et al., 2007), the intracellular loops extending from the MSDs interact with a groove in the NBDs. Thus, ATP-dependent changes in the association of the two NBDs cause a change in the position of the ICLs relative to the MSDs, inducing a conformational change in the MSDs. Thus, comprehension of inter-domain interactions is of paramount importance for the understanding of the molecular mechanism of CFTR activity. 4.1. NBD1–NBD2 interactions In CFTR, NBD1 interacts with NBD2 in the mode of a labile nucleotide sandwich, as depicted in Fig. 4A. This model is consistent not only with the prokaryotic MsbA and BtuCD entire transporters but also with several NBDs (for example, see Hung et al., 1998; Hopfner et al., 2000; Smith et al., 2002; Dawson and Locher, 2007; Ward et al., 2007). All the NBD structures reveal the same basic “tailto-head” fold, with highly conserved sequence motifs positioned to interact with bound ATP (Fig. 4A). This peculiar conformation of the CFTR–NBD dimer confers two nucleotide binding sites at the dimer interface, but only one consensus catalytic site. In the ‘NBD2 composite’ site, NBD2 contributes its nucleotide-binding residues

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of the Walker-A and Walker-B motifs and the H-loop, and NBD1 contributes its LSGGQ signature motif. The other interfacial site, the ‘NBD1 composite’ site, includes non-canonical residues from NBD1 Walker B (serine instead of glutamate) and switch (serine instead of histidine) motifs, as well as from the NBD2 signature sequence (LSHGH instead of LSGGQ). Not surprisingly, evidence suggests that the NBD2 composite site is catalytically competent, while ATP hydrolysis in the NBD1 composite site is smaller, or even null (Kidd et al., 2004; Ko and Pedersen, 1995; Cheung and Deber, 2008). NBD1–NBD2 dimerisation is attained in the CFTR protein in the presence of ATP (Vergani et al., 2003, 2005; Mense et al., 2006). Although no data for NBD1–NBD2 interaction at atomic resolution is available yet, SAXS experiments on isolated NBDs in solution have provided direct experimental evidence for NBD1–NBD2 dimerisation (Galeno et al., 2011; Galfrè et al., 2012). In the absence of ATP, NBD1 and NBD2, by themselves, in solution are monomeric globular proteins with a gyration radius of ∼1.8 nm. Addition of ATP induces the formation of NBD1 homodimers, but does not cause NBD2 dimerisation (Galeno et al., 2011; Galfrè et al., 2012). Interestingly, NBD1 does not form homodimers when the protein construct includes the RE (Lewis et al., 2004, 2005, 2010; Thibodeau et al., 2005). However, when the RE is removed, NBD1 in the presence of ATP crystallizes as a dimer (Atwell et al., 2010). Conversely, the mixture of NBD1-NBD2 in solution forms heterodimers, independent from the presence of nucleotides (Galeno et al., 2011; Galfrè et al., 2012). In the absence of ATP, the NBD1–NBD2 heterodimer has a bi-lobular arrangement with a gyration radius of 2.8 nm (Fig. 4B). Addition of ATP produces compaction of the heterodimer, forming a globular particle with a gyration radius of 2.1 nm (Fig. 4C). Interestingly, the shape of the NBD1–NBD2 heterodimer in solution obtained from SAXS experiments clearly resembles the conformation of these subunits in the closed (ATP absent) and open (ATP present) conformations of the CFTR channel in homology models (Mornon et al., 2009; see Fig. 4B and C). Using an overlay binding assay, Wang et al. (2002) identified regions of the NBDs involved in dimerisation. The NBD1 segments 581–628 and 477–512 interacted with NBD2, and the NBD2 segments 1280–1303, 1340–1366 and 1391–1422 were recognized to interact with NBD1 (Wang et al., 2002). Cysteine substitution crosslinking experiments identified some specific residues involved in NBD1–NBD2 interaction (Mense et al., 2006). Based on the length of the cross-linker, residue pairs between the NBD1 head, containing the Walker motifs, and the NBD2 tail, containing the ABC signature sequence, A462–S1347 and S459–V1379 were predicted at a ˚ and pair S434–D1336 at a distance of ∼16 A; ˚ the distance ≤8 A, residue pair S605–A1374 in central regions of NBD1 and NBD2 were ˚ a final cross-link across the NBD2 compredicted to lie at ≤8 A; posite site, between the signature sequence in the NBD1 tail and the Walker A sequence in the NBD2 head, including residue pairs S549–S1248 and S459–A1374 were estimated ≤8 A˚ apart. Notably, no cross linking was found between the NBD1 head and the NBD2 head (or NBD2 centre), nor between the NBD2 tail and the NBD1 tail (or NBD1 centre), confirming the “head-to-tail” conformation (Mense et al., 2006). 4.2. NBDs–ICLs interface The crystal structure of the bacterial ABC transporter proteins solved so far, in particular that of the bacterial exporter Sav1866 (Dawson and Locher, 2007; Dawson et al., 2007), shows its NBDs to be in close contact with both MSDs. Molecular models of CFTR were constructed based on its homology to Sav1866 (Mornon et al., 2008; Serohijos et al., 2008). These models predict inter-domain interactions between ICL2 and NBD2 and between ICL4 and NBD1.

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Interactions occur in a groove of the NBD1–NBD2 heterodimer especially rich in aromatic residues (Mornon et al., 2008; Serohijos et al., 2008). The other two intracellular loops, ICL1 and ICL3 interact with the NBD–ATP binding sites (Mornon et al., 2008). Cross-linking experiments have confirmed the proximity of the MSDs with their opposite NBDs through the ICLs: ICL1 and ICL2 with NBD2, and ICL3 and ICL4 with NBD1 (Serohijos et al., 2008; He et al., 2008). These data also showed that cross linking between domain-swapping interfaces at the ICLs and NBDs perturbs CFTR channel gating, suggesting that these interfaces are significantly involved in the stabilization of inter-domain contacts and regulation of channel gating. Two mutations in ICL4, L1065P and R1066C, prevent correct CFTR maturation (Seibert et al., 1996; Cotten et al., 1996). It is suggested that phenylalanine 508, the site of the most common CF mutation, F508del, which also impairs CFTR maturation, has a close interaction with residues at ICL4 (F1068, G1069) (Serohijos et al., 2008). This coincidence has prompted the hypothesis that this region is of key importance for CFTR folding (Serohijos et al., 2008). 4.3. NBDs–RD interactions and phosphorylation RD phosphorylation activates the CFTR channel by promoting the dimerization of NBD1–NBD2 (Mense et al., 2006). In fact, a direct interaction of the RD with NBDs was revealed by an overlay binding assay (Wang et al., 2002). RD-peptides bind to NBD1 segments between residues 514–535 and 544–580, and to NBD2 segments between 1271–1300, 1314–1356 and 1375–1411. Identification of the native and phosphorylated RD residues involved in the interaction with NBD1 and NBD2 were determined from NMR experiments (Baker et al., 2007; Bozoky et al., 2013). Multiple RD segments are involved in the interactions between the nonphosphorylated RD and NBDs, including the major interaction site surrounding S768. The majority of these interaction sites include the phosphorylation sites at 660, 700, 737, 795, and 813. The interaction of residues 661–671 with NBD1 is consistent with the crystal structures of NBD1, where the RE is in contact with the NBD1 core (Lewis et al., 2004, 2005; Thibodeau et al., 2005). Phosphorylation of the RD abolishes most binding to NBD1, with the exception of the region around S768 (Baker et al., 2007), and most biding to NBD2 (Bozoky et al., 2013).

5. Concluding remarks Study of the molecular structure of CFTR determines correct comprehension of the protein’s physiology, as well as understanding how perturbations introduced by mutations cause CF. Significant advancement towards the description of the CFTR structure has been achieved in the last ten years. The complexity of this protein has limited its study, and only an incomplete image of the CFTR molecular structure is now available. However, structural studies have informed the molecular mechanism of CFTR potentiators (Moran et al., 2005; Zegarra-Moran et al., 2007) and blockers (Dalton et al., 2012; Norimatsu et al., 2012) and have helped to identify CFTR correctors (Kalid et al., 2010; Odolczyk et al., 2013; He et al., 2013). Acquisition of complete structural information will guide better understanding of CFTR function and disease mechanisms. This knowledge will also permit the correct design of possible CF therapies.

Acknowledgements Thanks to Paolo Tammaro, Debora Baroni and Olga ZegarraMoran for comments and suggestions. This work was partially

supported by the Fondazione per la Ricerca sulla Fibrosi Cistica (Italy).

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