Facilitating Structure-Function Studies of CFTR Modulator Sites with Efficiencies in Mutagenesis and Functional Screening
The cystic fibrosis transmembrane conductance regulator (CFTR/ABCC7) is an ATP-dependent and phosphorylation-regulated plasma membrane chloride channel. It is composed of two membrane-spanning domains (MSDs), each containing six transmembrane helices, two intracellular nucleotide-binding domains (NBDs), and a regulatory (R) domain. The transmembrane helices from MSD1 and MSD2 form the channel pore, while the NBDs form the catalytic dimer that promotes channel gating, and the R domain regulates CFTR gating. Mutations in CFTR cause the genetic disease cystic fibrosis (CF), with the most common disease-causing mutations being the misprocessed p.Phe508del (present on one or both alleles in 90% of cases) and the gating mutation p.Gly551Asp (approximately 3%–4% of cases). Recently, a small-molecule therapy, Ivacaftor (Kalydeco or VX-770), developed by Vertex Pharmaceuticals, has been approved by the Food and Drug Administration for patients bearing the p.Gly551Asp mutation and several other less common CFTR gating mutations. VX-770 is a CFTR-specific potentiator capable of activating gating-incompetent channels in the clinical setting. However, despite this advancement, there remains no targeted therapy for the remaining approximately 95% of CF patients.
To address this unmet need, there is a requirement to assess other CFTR mutations for possible rescue by Kalydeco and to define the binding sites of such modulators on CFTR to enhance understanding of their mechanisms of action. This study describes a simple and rapid one-step PCR-based site-directed mutagenesis method to generate mutations in the CFTR gene. Using this method, CFTR mutants bearing deletions (p.Gln2_Trp846del, p.Ser700_Asp835del, p.Ile1234_Arg1239del) and truncations with polyhistidine tag insertion (p.Glu1172-3Gly-6-His*) were generated. These mutants either recapitulate disease phenotypes or serve as tools for modulator binding site identification. The drug responses of these mutants were evaluated using a high-throughput (384-well) membrane potential–sensitive fluorescence assay of CFTR channel activity within one week. This proof-of-concept study demonstrates that these methods enable rapid and quantitative comparison of multiple CFTR mutants to emerging drugs, facilitating future large-scale efforts to stratify mutants according to their “theratype” or most promising targeted therapy.
Approximately 2000 CFTR mutations have been identified to date, but molecular characterization has been performed on fewer than 10%. There is a pressing need to generate these uncharacterized CFTR mutations for in vitro study to test their response to emerging therapeutics and to discover targeted therapies that enhance the functional expression of CF disease-causing mutations. This approach could lead to the development of therapies tailored to each genotype rather than the current one-size-fits-all paradigm, which uses pharmacologic potentiators to treat multiple mutations affecting channel gating or corrector compounds targeting all mutations leading to CFTR misprocessing.
Site-directed mutagenesis has been fundamental in molecular biology and genetic research, allowing the generation of nucleotide substitutions to replicate disease-causing mutations in artificial expression systems to understand the molecular consequences leading to pathophysiology. Polymerase chain reaction (PCR)–based site-directed mutagenesis has been instrumental in analyzing the structure-function relationships of numerous disease-causing mutant proteins. Traditionally, this is done in double-stranded plasmid DNA containing the complementary DNA (cDNA) of the gene of interest. Various PCR approaches exist for site-directed cDNA mutations, including overlap extension PCR, inverted PCR, megaprimer PCR, and recombination PCR. The QuikChange kit by Stratagene has been the preferred method for over two decades for generating engineered mutations; however, it is not capable of performing deletion and insertion mutagenesis because it is optimized for single- or multiple-site primer-based modifications. Furthermore, insertion/deletion mutagenesis is incompatible with one-step QuikChange and other one-step PCR-based kits due to the relatively high error rate of their DNA polymerases, typically Taq polymerase, which lacks proofreading activity. To reduce second-site errors, molecular biologists often perform site-directed mutagenesis in smaller gene fragments and subsequently subclone the mutated fragment into an unmodified full-length template plasmid.
Using site-directed mutagenesis methods, the authors have generated over 100 CFTR missense mutant constructs, several of which have been recently reported and are critical tools in ongoing projects. However, many CF-associated mutations predicted to cause deletions, insertions, or truncations (stop mutations) remain to be generated, a process that is time-consuming. For example, a CF-associated mutation common in the Middle East (c.3700 A>G) leads to deletion of six amino acids, p.Ile1234_Arg1239del-CFTR, through alternative splicing. There is also a subset of at least 40 disease-causing mutations that fully or partially truncate NBD2 (residues 1173–1480), including various deletions and stop mutations. Future studies of the molecular defects caused by these mutations and assessment of their response to emerging therapies require rapid and efficient methods for mutagenesis and functional assessment, ideally in a multiwell high-throughput format.
To assess functional responses of CFTR variants in a high-throughput manner, the authors employed a rapid and simple mix-and-read fluorometric imaging plate reader (FLIPR) membrane potential assay. This method offers kinetic resolution comparable to electrophysiological measurements, allowing detection of rapid changes in membrane potential, but is faster, less labor-intensive, and higher throughput. FLIPR eliminates wash steps, resulting in healthier cells and shorter read times due to its simple mix-and-read protocol. Other membrane potential–sensitive dyes exist, such as DiBAC, but they have slower response times and are sensitive to temperature variations, making FLIPR more robust and yielding high signal-to-noise ratios. Importantly, FLIPR can measure both activation and inhibition of CFTR, whereas fluorescence-based halide dequenching assays detect only anion channel activation. A potential disadvantage is that non-CFTR ion channels may affect changes in membrane potential, but specificity can be assessed by confirming signature features of CFTR, including activation by cyclic AMP agonists, sensitivity to electrochemical anion gradients, and inhibition by CFTRinh-172.
In this study, an improved PCR-based site-directed insertion and deletion mutagenesis method was developed to generate disease-causing mutations in CFTR cDNA. This technique is rapid, inexpensive, and simple and can be used for replicating large vectors (>10 kb) without the need for subcloning. The authors incorporated desired insertions into primers (or omitted them for deletions) and used the KAPA HiFi HotStart PCR Kit, which contains a high-fidelity engineered B-family proofreading DNA polymerase, to generate four relatively complex CFTR mutants in a one-step PCR process. The deletion constructs removed 18 nucleotides (in-frame deletion of six amino acids in NBD2), rendering p.Ile1234_Arg1239del-CFTR; 405 nucleotides (in-frame deletion of residues 700–835 comprising the R domain), rendering p.Ser700_Asp835del-CFTR; and 2532 nucleotides (in-frame deletion of residues 2–846 comprising the MSD1-NBD1-R domain sequence), rendering p.Gln2_Trp846del-CFTR (an MSD2-NBD2 construct). The insertion construct added 27 nucleotides coding for two glycine residues, six histidine residues, and one stop codon, yielding p.Glu1172-3Gly-6His*-CFTR, a deletion mutant lacking 308 C-terminal residues (amino acids 1173–1480) and containing an engineered C-terminal poly-His tag. When transiently expressed in human embryonic kidney (HEK)-293 GripTite cells and paired with the rapid, multiwell (384-well) FLIPR assay of channel function, these methods permitted comparison of the functional consequences of emerging CFTR therapies on these rare deletion mutants.
The generation of mutant CFTR constructs was performed using the KAPA HiFi HotStart PCR Kit according to the manufacturer’s protocol with high-quality plasmid DNA containing wild-type CFTR cDNA (in pcDNA3.1) as the template. Specific PCR primers were designed for each mutant. For p.Ile1234_Arg1239del-CFTR, primers deleted 18 nucleotides; for p.Ser700_Asp835del-CFTR, 405 nucleotides were deleted; for p.Gln2_Trp846del-CFTR, 2532 nucleotides were deleted; and for p.Glu1172-3Gly-6His*-CFTR, 27 nucleotides were inserted encoding the glycine and histidine residues plus a stop codon. The PCR amplification used 1 or 10 ng of template plasmid and 0.3 µM final concentration of primers.
These methods enable efficient generation of complex CFTR mutations, facilitating rapid functional screening and aiding in the development of personalized therapies for cystic fibrosis patients with rare or complex mutations.
For PCR amplification, 1 or 10 ng of template plasmid and 0.3 µM (final concentration) of primers were used with the KAPA HiFi HotStart PCR Kit, following the manufacturer’s standard protocol. The PCR cycling conditions were optimized to ensure high-fidelity replication of the large CFTR cDNA plasmid (>10 kb) without the need for subcloning. The primers were designed to incorporate the desired deletions or insertions directly, enabling one-step generation of complex CFTR mutants.
The p.Ile1234_Arg1239del-CFTR mutant involved an in-frame deletion of 18 nucleotides corresponding to six amino acids in the nucleotide-binding domain 2 (NBD2). The p.Ser700_Asp835del-CFTR mutant contained a larger in-frame deletion of 405 nucleotides, removing residues 700 to 835, which encompass part of the regulatory (R) domain. The p.Gln2_Trp846del-CFTR mutant represented a substantial in-frame deletion of 2532 nucleotides, removing residues 2 to 846, which include membrane-spanning domain 1 (MSD1), NBD1, and the R domain, effectively creating a construct containing only MSD2 and NBD2. The p.Glu1172-3Gly-6His*-CFTR insertion mutant introduced 27 nucleotides encoding two glycine residues, six histidine residues (forming a polyhistidine tag), and a stop codon, resulting in a truncated CFTR protein lacking the C-terminal 308 amino acids (residues 1173–1480) with an engineered C-terminal His-tag for purification or detection purposes.
Following PCR amplification, the products were treated with DpnI restriction enzyme to digest the methylated, parental plasmid DNA template, enriching for the newly synthesized mutant DNA. The resulting DNA was then transformed into competent Escherichia coli cells for propagation and plasmid isolation. The presence of the intended mutations was confirmed by DNA sequencing.
Transient transfection of the mutant CFTR constructs into human embryonic kidney (HEK)-293 GripTite cells was performed to assess protein expression and function. The HEK-293 cells were cultured under standard conditions and transfected using established protocols optimized for high efficiency.
Functional analysis of the mutant CFTR proteins was conducted using a high-throughput, 384-well format membrane potential assay employing the FLIPR membrane potential–sensitive dye system. This assay allows rapid and quantitative measurement of CFTR chloride channel activity by detecting changes in membrane potential upon channel activation or inhibition. The assay was performed within one week of mutant generation, demonstrating the efficiency of the combined mutagenesis and functional screening approach.
The FLIPR assay was validated by confirming the signature features of CFTR channel activity, including activation by cyclic AMP agonists, sensitivity to electrochemical anion gradients, and inhibition by the specific CFTR inhibitor CFTRinh-172. This ensured that the measured responses were specific to CFTR function.
This integrated methodology of rapid one-step PCR-based mutagenesis combined with high-throughput functional screening facilitates the study of rare and complex CFTR mutations. It enables the stratification of CFTR mutants according to their response to emerging therapeutics, such as potentiators and correctors, thereby advancing personalized medicine approaches in cystic fibrosis.
In conclusion, the developed mutagenesis and screening platform provides a powerful tool for accelerating structure-function studies of CFTR modulator binding sites and assessing the theratype of diverse CFTR mutations. This approach supports ongoing efforts to expand targeted therapies beyond the small subset of mutations currently treatable with drugs like Ivacaftor, ultimately aiming to benefit the broader cystic ABBV-2222 fibrosis patient population.