The cystic fibrosis transmembrane conductance regulator (CFTR) anion channel is essential to maintain fluid homeostasis in key organs. Functional impairment of CFTR due to mutations in the cftr gene leads to cystic fibrosis. Here, we show that the first nucleotide-binding domain (NBD1) of CFTR can spontaneously adopt an alternate conformation that departs from the canonical NBD fold previously observed. Crystallography reveals that this conformation involves a topological reorganization of NBD1. Single-molecule fluorescence resonance energy transfer microscopy shows that the equilibrium between the conformations is regulated by adenosine triphosphate binding. However, under destabilizing conditions, such as the disease-causing mutation F508del, this conformational flexibility enables unfolding of the β-subdomain. Our data indicate that, in wild-type CFTR, this conformational transition of NBD1 regulates channel function, but, in the presence of the F508del mutation, it allows domain misfolding and subsequent protein degradation. Our work provides a framework to design conformation-specific therapeutics to prevent noxious transitions.
Subscribe to Journal
Get full journal access for 1 year
only $9.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Data supporting the findings of this manuscript are available from the corresponding author upon reasonable request. The atomic coordinates and structure factors reported in this paper have been deposited in the Protein Data Bank (PDB). The accession number for the structure reported in this paper is PDB 6ZE1. Source data are provided with this paper.
Riordan, J. R. et al. Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 245, 1066–1073 (1989).
Ratjen, F. et al. Cystic fibrosis. Nat. Rev. Dis. Primers 1, 15010 (2015).
Zhang, Z., Liu, F. & Chen, J. Molecular structure of the ATP-bound, phosphorylated human CFTR. Proc. Natl Acad. Sci. USA 115, 12757–12762 (2018).
Lukacs, G. L. & Verkman, A. S. CFTR: folding, misfolding and correcting the ΔF508 conformational defect. Trends Mol. Med. 18, 81–91 (2012).
Serohijos, A. W. R. et al. Phenylalanine-508 mediates a cytoplasmic-membrane domain contact in the CFTR 3D structure crucial to assembly and channel function. Proc. Natl Acad. Sci. USA 105, 3256–3261 (2008).
Protasevich, I. et al. Thermal unfolding studies show the disease causing ΔF508 mutation in CFTR thermodynamically destabilizes nucleotide-binding domain 1. Protein Sci. 19, 1917–1931 (2010).
Wang, C. et al. Integrated biophysical studies implicate partial unfolding of NBD1 of CFTR in the molecular pathogenesis of F508del cystic fibrosis. Protein Sci. 19, 1932–1947 (2010).
He, L. et al. Restoration of NBD1 thermal stability is necessary and sufficient to correct ΔF508 CFTR folding and assembly. J. Mol. Biol. 427, 106–120 (2015).
Rabeh, W. M. et al. Correction of both NBD1 energetics and domain interface is required to restore ΔF508 CFTR folding and function. Cell 148, 150–163 (2012).
Sharma, M., Benharouga, M., Hu, W. & Lukacs, G. L. Conformational and temperature-sensitive stability defects of the ΔF508 cystic fibrosis transmembrane conductance regulator in post-endoplasmic reticulum compartments. J. Biol. Chem. 276, 8942–8950 (2001).
Okiyoneda, T. et al. Peripheral protein quality control removes unfolded CFTR from the plasma membrane. Science 329, 805–810 (2010).
Lewis, H. A. et al. Impact of the ΔF508 mutation in first nucleotide-binding domain of human cystic fibrosis transmembrane conductance regulator on domain folding and structure. J. Biol. Chem. 280, 1346–1353 (2005).
Lewis, H. A. et al. Structure and dynamics of NBD1 from CFTR characterized using crystallography and hydrogen/deuterium exchange mass spectrometry. J. Mol. Biol. 396, 406–430 (2010).
Yang, Z. et al. Structural stability of purified human CFTR is systematically improved by mutations in nucleotide binding domain 1. Biochim. Biophys. Acta Biomembr. 1860, 1193–1204 (2018).
Aleksandrov, A. A. et al. Allosteric modulation balances thermodynamic stability and restores function of ΔF508 CFTR. J. Mol. Biol. 419, 41–60 (2012).
deCarvalho, A. C. V., Gansheroff, L. J. & Teem, J. L. Mutations in the nucleotide binding domain 1 signature motif region rescue processing and functional defects of cystic fibrosis transmembrane conductance regulator ΔF508. J. Biol. Chem. 277, 35896–35905 (2002).
Lewis, H. A. et al. Structure of nucleotide-binding domain 1 of the cystic fibrosis transmembrane conductance regulator. EMBO J. 23, 282–293 (2004).
Atwell, S. et al. Structures of a minimal human CFTR first nucleotide-binding domain as a monomer, head-to-tail homodimer, and pathogenic mutant. Protein Eng. Des. Sel. 23, 375–384 (2010).
Zhang, Z. & Chen, J. Atomic structure of the cystic fibrosis transmembrane conductance regulator. Cell 167, 1586–1597 (2016).
Liu, F., Zhang, Z., Csanády, L., Gadsby, D. C. & Chen, J. Molecular structure of the human CFTR ion channel. Cell 169, 85–95 (2017).
Aleksandrov, A. A. et al. Regulatory insertion removal restores maturation, stability and function of ΔF508 CFTR. J. Mol. Biol. 401, 194–210 (2010).
Lukacs, G. L. et al. Conformational maturation of CFTR but not its mutant counterpart (ΔF508) occurs in the endoplasmic reticulum and requires ATP. EMBO J. 13, 6076–6086 (1994).
Gong, X. et al. Non-native conformers of cystic fibrosis transmembrane conductance regulator NBD1 are recognized by Hsp27 and conjugated to SUMO-2 for degradation. J. Biol. Chem. 291, 2004–2017 (2016).
Aleksandrov, L., Aleksandrov, A. A., Chang, X. B. & Riordan, J. R. The first nucleotide binding domain of cystic fibrosis transmembrane conductance regulator is a site of stable nucleotide interaction, whereas the second is a site of rapid turnover. J. Biol. Chem. 277, 15419–15425 (2002).
Vergani, P., Lockless, S. W., Nairn, A. C. & Gadsby, D. C. CFTR channel opening by ATP-driven tight dimerization of its nucleotide-binding domains. Nature 433, 876–880 (2005).
Cui, L. et al. The role of cystic fibrosis transmembrane conductance regulator phenylalanine 508 side chain in ion channel gating. J. Physiol. 572, 347–358 (2006).
Mense, M. et al. In vivo phosphorylation of CFTR promotes formation of a nucleotide-binding domain heterodimer. EMBO J. 25, 4728–4739 (2006).
Kalinin, S., Valeri, A., Antonik, M., Felekyan, S. & Seidel, C. A. M. Detection of structural dynamics by FRET: a photon distribution and fluorescence lifetime analysis of systems with multiple states. J. Phys. Chem. B 114, 7983–7995 (2010).
Sigoillot, M. et al. Domain-interface dynamics of CFTR revealed by stabilizing nanobodies. Nat. Commun. 10, 2636 (2019).
Kalinin, S. et al. A toolkit and benchmark study for FRET-restrained high-precision structural modeling. Nat. Methods 9, 1218–1225 (2012).
Premchandar, A. et al. New insights into interactions between the nucleotide-binding domain of CFTR and keratin 8. Protein Sci. 26, 343–354 (2017).
Denning, G. M. et al. Processing of mutant cystic fibrosis transmembrane conductance regulator is temperature-sensitive. Nature 358, 761–764 (1992).
Proctor, E. A. et al. Rational coupled dynamics network manipulation rescues disease-relevant mutant cystic fibrosis transmembrane conductance regulator. Chem. Sci. 6, 1237–1246 (2015).
Kanelis, V., Hudson, R. P., Thibodeau, P. H., Thomas, P. J. & Forman-Kay, J. D. NMR evidence for differential phosphorylation-dependent interactions in WT and ΔF508 CFTR. EMBO J. 29, 263–277 (2010).
Csanády, L., Chan, K. W., Nairn, A. C. & Gadsby, D. C. Functional roles of nonconserved structural segments in CFTR’s NH2-terminal nucleotide binding domain. J. Gen. Physiol. 125, 43–55 (2005).
Wright, P. E. & Dyson, H. J. Intrinsically disordered proteins in cellular signalling and regulation. Nat. Rev. Mol. Cell Biol. 16, 18–29 (2015).
Ainscow, E. K., Mirshamsi, S., Tang, T., Ashford, M. L. J. & Rutter, G. A. Dynamic imaging of free cytosolic ATP concentration during fuel sensing by rat hypothalamic neurones: evidence for ATP-independent control of ATP-sensitive K+ channels. J. Physiol. 544, 429–445 (2002).
Cui, L. et al. Domain interdependence in the biosynthetic assembly of CFTR. J. Mol. Biol. 365, 981–994 (2007).
Goddard, T. D. et al. UCSF ChimeraX: meeting modern challenges in visualization and analysis. Protein Sci. 27, 14–25 (2018).
Li, S. J. & Hochstrasser, M. A new protease required for cell-cycle progression in yeast. Nature 398, 246–251 (1999).
Pardon, E. et al. A general protocol for the generation of nanobodies for structural biology. Nat. Protoc. 9, 674–693 (2014).
Vonrhein, C. et al. Data processing and analysis with the autoPROC toolbox. Acta Crystallogr. D Biol. Crystallogr. 67, 293–302 (2011).
Vagin, A. & Teplyakov, A. MOLREP: an automated program for molecular replacement. J. Appl. Crystallogr. 30, 1022–1025 (1997).
Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).
Bricogne, G. et al. BUSTER Version 2.10.1. Global Phasing Ltd. (2017).
Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D Biol. Crystallogr. 66, 12–21 (2010).
Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).
Hildebrandt, E. et al. A stable human-cell system overexpressing cystic fibrosis transmembrane conductance regulator recombinant protein at the cell surface. Mol. Biotechnol. 57, 391–405 (2015).
Sheppard, D. N. & Robinson, K. A. Mechanism of glibenclamide inhibition of cystic fibrosis transmembrane conductance regulator Cl− channels expressed in a murine cell line. J. Physiol. 503, 333–346 (1997).
Cai, Z., Taddei, A. & Sheppard, D. N. Differential sensitivity of the cystic fibrosis (CF)-associated mutants G551D and G1349D to potentiators of the cystic fibrosis transmembrane conductance regulator (CFTR) Cl− channel. J. Biol. Chem. 281, 1970–1977 (2006).
Rodrat, M. et al. Carbon monoxide-releasing molecules inhibit the cystic fibrosis transmembrane conductance regulator Cl− channel. Am. J. Physiol. Lung Cell. Mol. Physiol. 319, L997–L1009 (2020).
Wales, T. E., Eggertson, M. J. & Engen, J. R. Considerations in the analysis of hydrogen exchange mass spectrometry data. Methods Mol. Biol. 1007, 263–288 (2013).
Lau, A. M. C., Ahdash, Z., Martens, C. & Politis, A. Deuteros: software for rapid analysis and visualization of data from differential hydrogen deuterium exchange-mass spectrometry. Bioinformatics 35, 3171–3173 (2019).
Liu, H. & Naismith, J. H. An efficient one-step site-directed deletion, insertion, single and multiple-site plasmid mutagenesis protocol. BMC Biotechnol. 8, 91 (2008).
Talavera, A. et al. Phosphorylation decelerates conformational dynamics in bacterial translation elongation factors. Sci. Adv. 4, eaap9714 (2018).
Schrimpf, W., Barth, A., Hendrix, J. & Lamb, D. C. PAM: a framework for integrated analysis of imaging, single-molecule and ensemble fluorescence data. Biophys. J. 114, 1518–1528 (2018).
Hellenkamp, B. et al. Precision and accuracy of single-molecule FRET measurements—a multi-laboratory benchmark study. Nat. Methods 15, 669–676 (2018).
Kudryavtsev, V. et al. Combining MFD and PIE for accurate single-pair Förster resonance energy transfer measurements. ChemPhysChem 13, 1060–1078 (2012).
Tomov, T. E. et al. Disentangling subpopulations in single-molecule FRET and ALEX experiments with photon distribution analysis. Biophys. J. 102, 1163–1173 (2012).
Antonik, M., Felekyan, S., Gaiduk, A. & Seidel, C. A. M. Separating structural heterogeneities from stochastic variations in fluorescence resonance energy transfer distributions via photon distribution analysis. J. Phys. Chem. B 110, 6970–6978 (2006).
We thank C. R. O’Riordan for C127 cells, J. Kappes for HEK293 CFTR cells and J. R. Riordan, T. Jensen and CF Foundation Therapeutics for anti-CFTR antibodies. C.G. acknowledges support by the Fonds Forton, Welbio (grant no. CR-2012S-04R), Vaincre la Mucoviscidose, Mukoviszidose e.V., the Association Luxembourgeoise de Lutte contre la Mucoviscidose, ABCF2, Chiesi Fondation, the Cystic Fibrosis Foundation, Fondation Air Liquide and Fondation ULB. D.N.S acknowledges support from the CF Trust and CF Foundation Therapeutics. J.S. acknowledges support from Instruct-ERIC, part of the European Strategy Forum on Research Infrastructures (ESFRI), Instruct-ULTRA (EU H2020 grant no. 731005) and the Research Foundation—Flanders (FWO) for support with nanobody discovery. C.G. is a senior Research Associate of the FRS-FNRS. D.S. was a fellow of the FRIA. We acknowledge Diamond Light Source for time on beamlines i02, i04 and i24 under proposals 12718 and 9426. We are grateful to H. Remaut and B. Kobilka for careful reading of the manuscript. We thank F. Sobott and J. Ault of the Biomolecular Mass Spectrometry department in the Astbury Centre for their support and assistance in this work, as well as the BBSRC (BB/M012573/1) for funding.
The authors declare no competing interests.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
a, Representation of NBD1 containing two cysteines residues (yellow spheres) which are labeled by the fluorophores ATTO488 and Alexa647. b, FRET between both fluorophores attached to NBD1 is monitored while NBD1 diffuses through the confocal volume. c, Schematic illustration of the confocal microscope setup used here to study molecules freely diffusing in solution by multiparameter fluorescence detection with pulsed interleaved excitation (MFD-PIE)57,58. The lasers emit pulses with a fixed repetition rate and are synchronized by the TCSPC hardware (see below). An electronic delay leads to a lag time between green and red laser leading to interleaved excitation of the sample. Dichroic mirrors (DM) combine the lasers. They are then collimated and their beam profile is cleaned up by a single-mode optical fiber (SMF). Polychroic mirrors (PM) separate the excitation from the emission light both spectrally and spatially. The emission light is focused through a confocal pinhole and collimated by achromatic lenses (AL) to reduce chromatic aberrations. The emission light is then split up into two spectral ranges (green and red). Polarization beam splitters (PBS) further split the polarization of each color range into parallel (|) and antiparallel (⊥). Emission filters (EmF) finally transmit the appropriate spectral band and a lens focuses the emission light onto an avalanche photodiode (APD). The time-correlated single photon counting (TCSPC) hardware receives the signals from the APDs. Three parameters are registered per photon: the identity of the detector (red or green, parallel or antiparallel), the microtime (time since the last laser pulse) and the macrotime (time since the beginning of the experiment).
Extended Data Fig. 2 Acceptor fluorescence lifetime versus FRET efficiency plots for the 426–519 reporter pair.
a, Location of the smFRET reporters in the canonical conformation of NBD1 (based on the published structure of human NBD1, PDB: 2BBO). The positions of the labelled cysteines are shown as yellow spheres. The regulatory insertion (RI), which is not resolved in the structure, is depicted as a red dashed line. F508, located in the α-subdomain, and ATP are shown. b–d, Two-dimensional histograms of acceptor fluorescence lifetime (τA) versus FRET efficiency of the 426–519 reporter for the indicated conditions. The histograms provide a distinctive signature for each of the three observed FRET states. Small changes in acceptor lifetimes are due to an altered fluorescence quantum yield of the acceptor, likely due to sticking of the fluorophore to the protein. Correction of these effects did not substantially affect the observed FRET efficiency values (data not shown).
a, Differential scanning fluorescence (DSF) of purified NBD1 variants with and without nanobody G11a. The samples were incubated with SYPRO Orange dye and fluorescence was measured as a function of temperature. The melting temperatures (Tm) were determined by the maxima of the first derivative of fluorescence. Curves depict mean of duplicates of one experiment representative of at least three independent experiments. b, Binding of nanobody G11a to NBD1 variants measured by ELISA. Biotinylated NBD1 variants were immobilized on neutravidincoated plates and incubated with increasing concentrations of nanobody. Binding of nanobody was followed by immunodetection of the His6-tag (see Methods). Representative curve of 3 independent experiments is shown. Error bars represent the standard deviation (SD) of duplicates. See Supplementary Table 1 for pEC50 and Tm values.
FRET-restrained positioning and screening (FPS30) simulations of the accessible volumes of the donor and acceptor dyes ATTO488 and Alexa647 when attached to cysteine residues in NBD1. a, Accessible dye positions when attached to residues 426 and 519 in the β-strand swapped conformation observed in the 2PT:G11a structure. b, Accessible dye positions in the 479–519 reporter in the canonical conformation, based on the canonical NBD1 structure (PDB: 2BBO). Simulation results for all used reporters in both β-strand swapped and canonical conformations are detailed in Supplementary Table 3). Figures were generated using PyMOL (DeLano Scientific LLC).
a–c, Summary of patch-clamp studies of wild‐type CFTR Cl− channel in an excised inside‐out membrane patch from a C127 cell heterologously expressing wild‐type human CFTR. The data were acquired at 37 °C in the presence of ATP (0.3 mM) and PKA (75 nM) in the intracellular solution. After the channel was fully activated, G11a (1 μM) was then directly added to the intracellular solution bathing the membrane patch. A large Cl− concentration gradient was imposed across the membrane patch ([Cl−]int, 147 mM; [Cl−]ext, 10 mM) and membrane voltage was clamped at –50 mV. Single‐channel current amplitude (i) mean burst duration (MBD) and interburst interval (IBI) of wild‐type CFTR in the absence and presence of G11a are quantified. Symbols represent individual values and columns are means ± SEM (a, n = 6; b, c, n = 3); *, P = 0.03 vs control, Student’s t-test; two-sided. d–e, Open probability (Po) timecourses for an individual wild‐type CFTR Cl− channel in absence and presence of G11a (1 µM). Lowercase letters indicate the locations of the recordings shown in f and g during the Po timecourses. f-g, Representative recordings of an individual wild‐type CFTR Cl− channel in the absence and presence of G11a. The two recordings in the presence of G11a demonstrate the different gating patterns observed in its presence (b, steady‐state gating; c, high activity gating). Dotted lines indicate where the channel is closed and downward deflections correspond to channel openings.
a-b. Structures of the canonical conformation (PDB: 2BBO) and the β-strand swapped conformation reported here. Unresolved segments are depicted in dashed lines, the RI (404–436) in red and the N-terminal region (389–403) in magenta. c-d, Donor lifetime in presence of acceptor (τD(A)) vs FRET efficiency histograms of the 390–519 reporter pair with and without ATP. The red line is called the ‘static FRET line’ and characterizes the theoretical relationship between lifetime and FRET efficiency in absence of conformational dynamics on the timescale of the measurement (milliseconds). e, Overlay of FRET efficiency histograms of 2PT-390-519 without ATP and with 2 mM ATP. Raw data (fill) was fitted using the PDAFit software (thick lines, see Methods). f-g, Donor lifetime vs FRET efficiency plots of 2PT-390-519 without the RI and with the RI in presence of nanobody G11a. h, Reversibility of the conformational change was tested. At the beginning of the experiment no ATP was present and after 2500 s ATP was added to yield a final concentration of 2 mM. i, Deuterium build-up curves of the 392–399 peptide identified in mass spectral analyses of NBD1-2PT (blue) and ΔRI-NBD1 (black). Error bars represent the standard deviation of technical triplicates. Data shown are representative of three biological replicates.
Cells that expressed CFTR and were successfully permeabilized were selected by gating for EGFP + (left, laser: 488 nm – detector: 525/50 nm) and DAPI + (right, laser: 405 nm – detector: 450/50 nm) as shown for each of the three nanobodies tested: the negative control, T2a and G11a (as indicated).
Statistical source data.
Statistical source data.
Unprocessed western blot.
Statistical source data.
Statistical source data.
Statistical source data.
Statistical source data.
Statistical source data.
Statistical source data.
About this article
Cite this article
Scholl, D., Sigoillot, M., Overtus, M. et al. A topological switch in CFTR modulates channel activity and sensitivity to unfolding. Nat Chem Biol 17, 989–997 (2021). https://doi.org/10.1038/s41589-021-00844-0