Ribosomal frameshifting selectively modulates the assembly, function, and pharmacological rescue of a misfolded CFTR variant
Edited by Jonathan Weissman, Whitehead Institute, Cambridge, MA; received August 1, 2024; accepted August 19, 2024
Significance
Many diseases stem from imbalances between protein synthesis and degradation that arise from mutations and/or cellular stressors. The molecular mechanisms of such lapses in proteostasis often coincide with aberrant regulation of protein translation. Here, we identify a structure within the transcript encoding the cystic fibrosis transmembrane conductance regulator channel that allows the ribosome to halt translation in response to cotranslational misfolding. We show that this motif modifies the assembly, function, and pharmacological properties of the most common cystic fibrosis variant. This cross talk between the ribosome and nascent polypeptide allows the ribosome to adjust its activity to reduce the synthesis of misfolded proteins. These findings suggest that ribosomal frameshifting and premature translational termination play a role in protein quality control.
Abstract
The cotranslational misfolding of the cystic fibrosis transmembrane conductance regulator chloride channel (CFTR) plays a central role in the molecular basis of CF. The misfolding of the most common CF variant (ΔF508) remodels both the translational regulation and quality control of CFTR. Nevertheless, it is unclear how the misassembly of the nascent polypeptide may directly influence the activity of the translation machinery. In this work, we identify a structural motif within the CFTR transcript that stimulates efficient −1 ribosomal frameshifting and triggers the premature termination of translation. Though this motif does not appear to impact the interactome of wild-type CFTR, silent mutations that disrupt this RNA structure alter the association of nascent ΔF508 CFTR with numerous translation and quality control proteins. Moreover, disrupting this RNA structure enhances the functional gating of the ΔF508 CFTR channel at the plasma membrane and its pharmacological rescue by the CFTR modulators contained in the CF drug Trikafta. The effects of the RNA structure on ΔF508 CFTR appear to be attenuated in the absence of the ER membrane protein complex, which was previously found to modulate ribosome collisions during “preemptive quality control” of a misfolded CFTR homolog. Together, our results reveal that ribosomal frameshifting selectively modulates the assembly, function, and pharmacological rescue of a misfolded CFTR variant. These findings suggest that interactions between the nascent chain, quality control machinery, and ribosome may dynamically modulate ribosomal frameshifting in order to tune the processivity of translation in response to cotranslational misfolding.
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Cystic fibrosis (CF) is a disease of protein homeostasis (proteostasis) that arises from the defective biosynthesis, folding, and/or function of a chloride channel known as the cystic fibrosis transmembrane conductance regulator (CFTR) (1, 2). The most common CF mutation (ΔF508) induces CFTR misfolding by decoupling the folding of its subdomains during translation (3, 4). This cotranslational misassembly reaction enhances the retention and degradation of the CFTR protein within the endoplasmic reticulum (ER) and ultimately reduces the trafficking of the functional protein to the plasma membrane. This lapse in protein quality control (QC) coincides with the remodeling of the CFTR proteostasis network and changes in the dynamics of CFTR translation (5–11). Nevertheless, the precise chain of events involved in the cross talk between the conformational state of nascent CFTR, the activity of the translation machinery, and the interaction of the nascent chain with various components of the cellular proteostasis network is not fully understood.
CFTR QC begins during its cotranslational folding at the ribosome–translocon complex (12). Vectorial folding of its five subdomains is orchestrated, in part, by positional variations in translation kinetics that are coordinated by rare codons (9), the relative abundance of certain translation factors (5, 7), and the presence of certain messenger RNA (mRNA) secondary structures (13, 14). Together, these effectors help the ribosome tailor its activity to the kinetic constraints of cotranslational CFTR folding. Indeed, the effects of CF mutations on CFTR folding and assembly are highly sensitive to changes in translational dynamics. For instance, suppressing translation initiation reduces the density of ribosomes on the ΔF508 transcript in a manner that partially rescues the stability, cellular trafficking, and function of the ΔF508 CFTR protein (7, 11). The cotranslational misfolding of a ΔF508 CFTR homolog (ΔF670 Yor1) also appears to disrupt translation by enhancing ribosome collisions and triggering “preemptive quality control”—a cotranslational QC pathway mediated by the ER membrane protein complex (EMC) (11). Though the later finding suggests that the intrinsic activity of the ribosome is sensitive to conformational transitions in the nascent chain, it remains unclear whether cotranslational misfolding events are capable of directly altering translation in real time.
We recently found that cotranslational folding mechanically modulates a translational recoding mechanism known as −1 programmed ribosomal frameshifting (−1PRF) during viral polyprotein biosynthesis (15, 16). These findings reveal that under certain circumstances, the ribosome exhibits an enhanced propensity to slip into alternative reading frames in response to conformational transitions in the nascent polypeptide chain (17). In the following, we evaluate whether similar feedback occurs during CFTR biosynthesis. We first identify a structured segment within the region of the CFTR transcript that stimulates ribosomal frameshifting (RF). We then show that the disruption of this motif selectively remodels the interactome of ΔF508 CFTR in a manner that partially restores its expression and function. These modifications, which have minimal impact on WT CFTR, also enhance the pharmacological rescue of ΔF508 CFTR by one of the leading CF therapeutics known as Trikafta. Our results show that this feedback hinges on the EMC, suggesting that this mechanism is involved in preemptive QC. Our findings collectively suggest that this RF site acts as a QC-mediated translational “kill switch” that selectively promotes the premature termination of translation in response to the cotranslational misfolding of the nascent chain.
Results
Identification of an Active −1 Ribosomal Frameshift Site in the CFTR Transcript.
Our recent findings show that the mechanical forces generated by cotranslational folding can stimulate −1PRF (15, 16), which minimally requires a slippery sequence within the mRNA and an adjacent stem-loop (18). To identify potential RF sites in the CFTR transcript we first searched for consensus or near-consensus X1 XXY4 YYZ7 slippery sequences, where XXX and YYY are nucleobase triplets and Z can be any nucleobase (19–21). We identified five consensus X1 XXY4 YYZ7 sites and eight other near-consensus sites distributed throughout the transcript (Fig. 1A and SI Appendix, Fig. S1). Three of these sites are clustered within the region encoding nucleotide binding domain 2 (NBD2) and are proximal to two predicted RNA stem-loops (Fig. 1A). A uridine-rich sequence referred to here as slip-site A (UUU AUU UUU UCU, SSA) lies upstream of the canonical slip-site B (A1 AAA4 AAC7, SSB). SSB is eight nucleotides upstream of a small, predicted stem-loop (SL2). We also identified a third putative noncanonical slip-site (A1 GAA4 AUA7, SSC) downstream of SSB that is positioned seven nucleotides upstream of a much longer putative stem-loop (SL2); an ideal spacing for the stimulation of −1PRF (18).
Fig. 1.
Given that ribosome collisions can occur during CFTR translation (11) and that they are known to modulate −1PRF (22), we hypothesized that RF could occur at one or more of these sites. To determine whether efficient −1 RF occurs at any of these positions, we generated a series of bicistronic reporters in which a short segment of the CFTR transcript bearing these features is inserted between an upstream Renilla luciferase (RLuc, expression control) and a downstream −1 firefly luciferase (FLuc, −1 RF reporter, Fig. 2A) (23). We generated three versions of this reporter that produce fLuc in response to frameshifting at any of these slip-sites (SSABC), or conditionally at B or C (SSBC), or at C only (SSC) by knocking out various −1-frame stop codons downstream of these slip-sites (Fig. 2A orange). Transient expression of the SSABC reporter in a human bronchial epithelial cell line (CFBE41o-) generates a detectable FLuc frameshift signal fourfold over the no-insert control baseline. RLuc intensities are similar for all three reporters, suggesting that all three reporters exhibit similar expression under these conditions (Fig. 2B). FLuc intensities are also quite similar for all three reporters (Fig. 2C), which implies that most frameshifting occurs at SSC. Consistent with this interpretation, we find that the FLuc signal is ablated by mutations that disrupt SSC (SSCmut, Fig. 2C). We note that we ruled out splicing artifacts (24) by RT-PCR and found that the frameshift signal can also be partially suppressed by stop codons in either upstream of SSC in the 0-frame (5′Ter) or downstream of SSC in the −1-frame (3′Ter, Fig. 2D and SI Appendix, Fig. S2), which confirms that these signals arise from a genuine RF. By normalizing the RLuc and FLuc signals, we estimate ribosomal frameshifting occurs with an efficiency of 2.9 ± 0.2% at SSC, which is comparable to levels achieved during translation of the canonical HIV gag-pol motif (5.2 ± 0.4%, Fig. 2D). Similar frameshifting also occurs in HEK293T cells (SI Appendix, Fig. S3). Together, these results identify an active −1 RF motif within the human CFTR transcript.
Fig. 2.
Impact of Ribosomal Frameshifting on the CFTR Interactome.
Unlike viral −1PRF events that generate functionally distinct proteins, a −1 frameshift at SSC should only fuse five non-native residues onto nascent CFTR prior to translational termination and the truncation of the channel protein in the middle of NBD2 (Fig. 1A). Based on this consideration, we hypothesized that the RF motif serves to promote frameshifting and the premature termination of translation in response to cotranslational CFTR misassembly. To determine whether this motif is involved in assembly, we assessed how this RNA structure influences the interactomes of WT and ΔF508 CFTR. Briefly, we designed a series of mutations to disrupt the slip-sites and weaken the stem-loops within NBD2 while preserving the amino acid sequence of CFTR in the 0-frame (RFmut, Fig. 1B). We then incorporated these modifications into the full-length CFTR (Fig. 1C) and used affinity purification-mass spectrometry (AP-MS) based interactome profiling to determine the effects of these silent modifications on the interactions formed by WT and ΔF508 CFTR (5, 25, 26). In agreement with previous reports (5, 25), the ΔF508 mutation significantly enhances the interaction of CFTR with numerous proteostasis factors (Fig. 3A and SI Appendix, Table S1). By comparison, the incorporation of the silent mutations that disrupt the RNA structure in NBD2 have minimal impact on the WT CFTR interactome (WT-RFmut, Fig. 3A). In contrast, these same mutations significantly alter the propensity of ΔF508 CFTR to interact with several translation factors and quality control proteins (ΔF508-RFmut, Fig. 3A). Consistent with our hypothesis, the disparate effects of this transcript modification on WT and ΔF508 suggests that the native RNA structure in NBD2 selectively alters CFTR translation and assembly in response to misfolding. We note that the ΔF508 mutation is ~2.4 kb upstream of the ribosomal frameshift site in NBD2 (Fig. 1C), which suggests that these differences likely arise from the effects of the ΔF508 mutation on the nascent CFTR protein rather than any perturbation of the mRNA structure that may arise from the deletion of these three bases.
Fig. 3.
There are several key differences in the interaction profiles of ΔF508 and ΔF508-RFmut CFTR (SI Appendix, Table S1). We first note that about half of the aberrant interactions formed by ΔF508 CFTR are unaffected by these silent mutations (see Cluster 1 in Fig. 3B & Clusters 2 to 3 in SI Appendix, Figs. S4 and S5). As both genetic constructs encode the same protein in the 0-frame, we suspect these common interactions may arise from the posttranslational effects of the ΔF508 mutation on the CFTR protein. Nevertheless, ΔF508-RFmut exhibits significantly attenuated associations with eight ribosomal proteins (Fig. 3B and SI Appendix, Table S1). These silent mutations also appear to modulate QC, as ΔF508-RFmut exhibits both weaker interactions with certain chaperones and cochaperones (e.g., CANX, HspA6, BAG2, and DNAJB6) and stronger interactions with others (e.g., HspB1, HspA4, and DNAJA3 & SI Appendix, Fig. S5). These QC modifications also coincide with a decrease in the interaction of ΔF508-RFmut with the UBE3C ubiquitin ligase, the ER-PHAGY protein RTN3, and various components of the 19S proteasome complex (i.e., PSMC1, PSMC2, and PSMC4, SI Appendix, Table S1), which suggests that the RF motif ultimately modifies CFTR degradation. We note these silent mutations enhance the interaction of ΔF508 CFTR with two components of the eIF3 complex (i.e., eIF3c and eIF3e, Cluster 3, SI Appendix, Fig. S4 and Table S1), which is known to modify the proteostatic effects of the ΔF508 mutation (7). Finally, these silent mutations also attenuate interaction with a subunit of the eIF2 complex (eiF2B2), whose alpha subunit was recently seen to regulate −1PRF in SARS-CoV-2. Overall, these results suggest that the ribosomal frameshift site in NBD2 acts as a junction where cotranslational misfolding triggers changes in CFTR translation, QC, and degradation.
Impact of Ribosomal Frameshifting on CFTR Expression.
To determine how these changes in cotranslational assembly impact CFTR proteostasis, we compared the effects of the RFmut modification on ΔF508 and WT CFTR expression. Western blot analyses reveal that modifications of the RF motif have minimal impact on the maturation of WT or ΔF508 CFTR in both CFBE41o- and HEK293T cells as judged by the relative abundance of the mature (C band) and immature (B band) glycoforms (Fig. 4 A–C and SI Appendix, Fig. S6). This observation is perhaps unsurprising given that the RFmut modification does not ultimately change the amino acid sequence of the dominant CFTR translation products, which are still subject to many of the same posttranslational QC interactions (Fig. 3A). The RFmut modification also appears to have little, if any, impact on total CFTR levels in CFBE41o- cells, as detected by western blotting (Fig. 4D). Any subtle differences in expression under these conditions are likely masked by the large variations in the transcript levels generated by transient transfections (SI Appendix, Fig. S7). To quantitatively compare cellular CFTR accumulation, we used flow cytometry to compare single-cell surface and intracellular CFTR immunostaining levels in HEK293T cells (Materials and Methods). Flow cytometry measurements confirm that disrupting the RF motif does has no impact on the WT CFTR expression and does not rescue the plasma membrane expression of ΔF508 CFTR (Fig. 4 E and F). However, the RFmut modification does increase intracellular ΔF508 levels by 19 ± 11% (P = 0.007, Fig. 4 E and F). Notably, the intracellular immunostaining intensity of ΔF508-RFmut is indistinguishable from that of WT (Fig. 4E), which suggests that this motif normally contributes to the reduced accumulation of ΔF508 CFTR in the ER. While all trends in expression were generally consistent in CFBE41o- and HEK293T cells (Fig. 4 and SI Appendix, Fig. S6), this enhanced intracellular accumulation of ΔF508-RFmut CFTR was not observed in the context of CRISPR knockout cells lacking the EMC (SI Appendix, Figs. S8 and S9). This observed connection to the EMC potentially suggests that this feedback may play a role in preemptive QC. Though changes in expression are subtle, our results reveal that the RF motif selectively reduces the expression of ΔF508 CFTR in an EMC-dependent manner- potentially by promoting the premature termination of translation in response to misfolding.
Fig. 4.
Impact of Ribosomal Frameshifting on CFTR Function.
To determine how the RF motif impacts CFTR function, we compared the effects of the RFmut modification on the ΔF508 and WT CFTR-mediated quenching of a cellular halide-sensitive yellow fluorescent protein (hYFP) (27). Briefly, we generated a series of recombinant stable HEK293T cell lines that inducibly express a single CFTR variant off a bicistronic transcript containing a downstream internal ribosomal entry site (IRES) that produces the hYFP sensor and a fluorescent mKate reference fluorophore (Fig. 5A). We then used flow cytometry to track the CFTR-mediated quenching of hYFP fluorescence in relation to an mKate reference fluorophore at the single-cell upon addition of iodide (Fig. 5B). Cells expressing CFTR undergo an initial rapid hYFP quenching phase that occurs within the dead time for mixing that is followed by a slow exponential decay in the single-cell hYFP: mKate values (Fig. 5B). The observable hYFP quenching phase was ninefold slower, on average, among cells expressing ΔF508 CFTR and was further delayed by a CFTR-specific inhibitor (CFTR(Inh)-172, Fig. 5B and SI Appendix, Fig. S10), which suggests that the observed quenching is rate limited by CFTR channel conductance. Half-life values derived from a global fit of the slow hYFP quenching phase reveal that the RFmut modification has minimal impact on WT CFTR but decreases the half-life of ΔF508 CFTR-mediated hYFP quenching by 46 ± 6% (Fig. 5 B and C). These observations suggest that the mutagenic disruption of the RF motif and its resulting impact on the CFTR interactome partially restore the function of ΔF508 CFTR. Interestingly, we find that the disruption of the RF motif does not result in an uptick in ΔF508 CFTR conductance in the context of EMC knockout cells (SI Appendix, Fig. S11), which again suggests that this feedback is EMC mediated.
Fig. 5.
Disrupting the RF motif enhances hYFP quenching by ΔF508 CFTR without increasing its plasma membrane expression (Fig. 4F), which indicates that this modification may improve the function of mature ΔF508 CFTR. We therefore compared the channel open probability (Po) of these variants in the context of inside-out macro- and/or micropatches excised from the plasma membranes of transfected HEK293T cells. Briefly, we monitored single channel activity over time (8 to 13 min) in the presence of 1.5 mM ATP and Protein Kinase A (PKA, 124 U/mL) at a holding potential of 60 mV. We then estimated the number of channels in the patch and verified the identity of the channels with the sequential addition of a potentiator (VX-770) and a CFTR inhibitor [CFTR (inh)-172]. Fig. 5 D and E shows representative current traces for patches containing ΔF508 and ΔF508 CFTR RFmut CFTR, respectively. Currents across patches containing ΔF508-RFmut CFTR generally feature more channel opening events relative to those containing ΔF508 CFTR (Fig. 5 D and E). Overall, disruption of the RF motif increases the ΔF508 CFTR Po by 1.6-fold (P = 0.025, Fig. 5F). The RFmut modification has a similar impact on WT CFTR opening despite the fact that it has limited impact on the slow phase of cellular hYFP quenching (Fig. 5 C and F). Nevertheless, it is possible that the effect of the RFmut modification of the single channel behavior of WT CFTR may be more pronounced within the initial rapid quenching phase that occurs within the mixing time. Regardless, these results confirm that the resulting modifications to the QC of nascent ΔF508 CFTR in the ER enhance the activity of the mature channel at the plasma membrane. Together, these cumulative findings reveal that the RF motif selectively modulates the assembly of ΔF508 CFTR in a manner that ultimately attenuates its expression and function.
Impact of Ribosomal Frameshifting on the Pharmacological Rescue of ΔF508 CFTR.
The functional expression of ΔF508 and various other CF variants can be partially restored by Trikafta, an FDA-approved cocktail of two correctors that stabilize the CFTR protein (VX-661 + VX-445) and a potentiator that activates it (VX-770) (28–30). To determine whether ribosomal frameshifting potentially influences the effects of these compounds, we assessed whether the RF motif impacts the pharmacological rescue of ΔF508 CFTR. As expected, flow cytometry-based immunostaining measurements reveal that both WT and ΔF508 exhibit enhanced accumulation at the plasma membrane and within the secretory pathway upon stabilization by VX-661 + VX-445 (Fig. 6 A and B). A western blot analysis also confirms that these correctors enhance the maturation of both variants (Fig. 6C and SI Appendix, Fig. S12). However, similar gains in expression and maturation were observed for WT-RFmut and ΔF508-RFmut (Fig. 6 A–C and SI Appendix, Fig. S12), which demonstrates that the RF motif has minimal impact on ΔF508 expression upon stabilization by corrector molecules. Next, we assessed functional rescue of CFTR conductance using our hYFP quenching assay. For this purpose, the potentiator VX-770 was added in combination with the correctors VX-661 and VX-445 to evaluate the functional rescue by Trikafta. As a control, these quenching assays were also performed in the presence of a channel inhibitor (SI Appendix, Fig. S10). Despite the comparable expression of ΔF508 and ΔF508-RFmut in the presence of correctors, HEK293T cells expressing ΔF508-RFmut that were treated with VX-661 + VX-445 + VX-770 exhibit a is 41 ± 5% decrease in hYFP quenching half-life relative to cells expressing ΔF508 CFTR under the same conditions (Fig. 6D). Together, these results suggest that while the RF motif has little impact on the correction of ΔF508 CFTR expression, disrupting the RNA structure that promotes frameshifting ultimately enhances its functional rescue.
Fig. 6.
Discussion
The ΔF508 mutation promotes cotranslational CFTR misfolding and degradation in a manner that reduces its functional expression (1). Nevertheless, its proteostatic effects also stem from aberrant translational dynamics. It was previously shown that the impact of the ΔF508 mutation on the secondary structure of the CFTR transcript perturbs translation dynamics in a manner that compromises channel expression and function (13, 14). Indeed, this mutation modifies interactions with over 20 translation factors and RNA processing proteins involved in CFTR biosynthesis (5). Moreover, the cotranslational misfolding of a yeast ΔF508 CFTR homolog (ΔF670 Yor1) triggers interactions with the EMC in a manner that results in ribosome collisions and the premature termination of translation (11). However, it remains unclear how the cotranslational misfolding of the ΔF508 protein could modify the activity of the ribosome itself. Given that conformational transitions in the nascent chain can stimulate −1PRF (15, 16, 31), we searched the CFTR transcript for structural features that might promote ribosomal frameshifting. Our findings identified a slippery sequence (SSC) and stem-loop (SL2) that stimulate ribosomal frameshifting and the premature translational termination (Fig. 2). To determine how this RF motif impacts CFTR expression and function, we designed a series of silent mutations that maintain the native amino acid sequence in the 0-frame while disrupting these putative RNA structures (RFmut, Fig. 1). While this modification has minimal impact on WT CFTR assembly, disrupting the RF motif significantly remodels ΔF508 CFTR interactome (Fig. 3). These changes in ΔF508 assembly slightly increase the accumulation of ΔF508 CFTR within the secretory pathway but markedly enhance its functional conductance at the plasma membrane (Figs. 4 and 5). Finally, we show that disrupting the RF motif also enhances the pharmacological rescue of ΔF508 CFTR function (Fig. 6). Given that the disruption of this motif has minimal impact on the assembly, expression, or pharmacological profile of WT CFTR (Figs. 2–5), we conclude that this RNA structure stimulates RF and the premature termination of translation in response to cotranslational CFTR misfolding. Together, our findings suggest that the RF motif acts as a “kill switch” that down-regulates the synthesis of defective proteins. Future investigations are needed to determine how this translational regulation influences the biosynthesis and pharmacological response of other known CF variants.
Interactome profiles suggest that this RF motif may represent an assembly junction where the conformational state of the nascent chain modulates the interplay between translation and QC. We identify several ΔF508-specific chaperone interactions that are lost and others that are gained upon disruption of the RF motif (SI Appendix, Table S1 and Fig. 3). Moreover, weakening this RNA structure in the context of the ΔF508 transcript significantly reduces the coimmunoprecipitation of several ribosomal proteins (SI Appendix, Table S1 and Fig. 3), which implies that ribosomes typically stall within this region during ΔF508 translation. Based on these observations, we propose that the misassembly of nascent ΔF508 CFTR stimulates ribosomal frameshifting and directs the ribosome to a stop codon in the −1-frame (Fig. 7). Without the prompt intervention of ribosome quality control (32), this premature termination may cause trailing ribosomes to collide (Fig. 7). Such cross talk may explain the link between cotranslational misfolding and ribosome collisions that occurs during preemptive QC (11). This interpretation is consistent with our observations that the effects of the frameshift motif are attenuated in EMC knockout cells (SI Appendix, Figs. S9 and S11). Ribosomal pausing (11) and/or heightened frameshifting during ΔF508 CFTR translation could potentially explain how this mutation enhances associations with UPF1 and other nonsense-mediated decay (NMD) proteins (5) that typically promote the degradation of nonsense transcripts (33, 34). Knocking down UPF1 and other associated RNA binding proteins partially restores ΔF508 expression (5), which until now has been a perplexing observation given that ΔF508 preserves the native reading frame. The stimulation of frameshifting by ΔF508 would promote premature termination via stop codons in the −1-frame. Such a link between ribosomal frameshifting and NMD has been previously described (35), and certain ribosome-associated molecular chaperones have been found to influence −1PRF (36). Nevertheless, to our knowledge, previous investigations have not established a direct role of ribosomal frameshifting in protein QC. We note that these interpretations remain speculative given that we are unable to compare ribosomal frameshifting efficiencies during CFTR biosynthesis. We also note that our interactome measurements do not directly identify EMC interactions or reveal a clear link to NMD, which potentially suggests that these interactions may be weak or transient. Additional investigations are needed to gain mechanistic insights into the interplay between cotranslational misfolding, ribosomal frameshifting, and RNA surveillance pathways.
Fig. 7.
There are currently few examples of bona fide −1PRF sites within eukaryotic transcripts and the current evidence for such remains contentious (24, 37). One key line of evidence that has cast doubt on the role of ribosomal frameshifting in eukaryotic gene regulation involves the general lack of the extended coding sequences within the alternative reading frames of eukaryotic genes—a classic indication of translational recoding in viral transcripts. Due to an abundance of stop codons in the −1 reading frame, ribosomal frameshifting in eukaryotes should most often result in premature termination of translation- an outcome that ribosomes have generally evolved to avoid. Nevertheless, premature termination and/ or the destabilization of the transcript may be beneficial under conditions in which the nascent polypeptides begin to misfold. Cells routinely undergo swings in adaptive proteostasis where the products of translation and the pool of molecular chaperones they rely on must be remodeled on short time scales. Based on our findings, we propose that this putative connection between cotranslational misfolding, ribosomal frameshifting, and RNA surveillance pathways provides a means for cells to remodel the transcriptome in response to misassembly in order to prevent the accumulation of misfolded proteins. Additional investigations are needed to determine whether similar QC checkpoints are present in the transcripts of other proteins that are predisposed to misfolding. Such investigations may offer additional mechanistic insights into the molecular basis of a wide array of diseases of aberrant cellular proteostasis.
Materials and Methods
Cell Culture.
HEK293T cells were grown in Dulbecco’s modified Eagle’s medium (Gibco, Grand Island, NY) containing 10% fetal bovine serum (Corning, Corning, NY) and a penicillin (100 U/mL)/streptomycin (100 µg/mL) antibiotic supplement (Gibco, Grand Island, NY) in a humidified incubator containing 5% CO2 at 37 °C. HEK293T cells used for interactome measurements were grown under similar conditions but with 1% L-glutamine (200 mM) supplementation. Plasmid DNA constructs were transiently expressed in HEK293T cells using Lipofectamine 3000 (ThermoFisher Scientific, cat #L3000015). Cells were dosed with CFTR modulators a day after transfection when applicable. Two days posttransfection, cells were washed with 1× phosphate-buffered saline (AthenaES, Baltimore, MD) and harvested with 1× 0.25% Trypsin-EDTA (Gibco, Grand Island, NY).
CFBE41o- cells were grown in minimal essential medium (Gibco, Grand Island, NY) containing 10% fetal bovine serum (Corning, Corning, NY) and a penicillin (100 U/mL)/streptomycin (100 µg/mL) (Gibco, Grand Island, NY) antibiotic supplement (Gibco, Grand Island, NY) in a humidified incubator containing 5% CO2 at 37 °C on culture plates coated with PureCol purified bovine collagen (Advanced BioMatrix, Carlsbad, CA). CFBE41o- cells were transfected using Lipofectamine 3000 (ThermoFisher Scientific, cat #L3000015). Two days posttransfection, cells were washed with 1× HEPES-buffered saline (Gibco, Grand Island, NY) and harvested with 1× 0.25% Trypsin-EDTA (Gibco, Grand Island, NY).
CFTR function measurements were carried out in recombinant stable cells were made from genetically modified HEK293T cells grown in 10 cm dishes in complete media as was previously described (38, 39). Briefly these HEK293T cells contain a Tet-Bxb1-BFP “landing pad” where recombination results in the integration of a plasmid into the gDNA of the cell (38). These cells were cotransfected with plasmid and a Bxb1 recombinase expression vector using Fugene 6 (Promega, Madison, WI). Doxycycline (2 µg/mL) was added 1 d after transfection and the cells were grown for 3 d at 33 °C. The cells were then incubated at 37 °C for 24 h prior to passaging for further experiments.
Mass Spectrometry–Based Interactome Profiling.
A pcDNA5 vector containing untagged CFTR with a CMV promoter was used in the interactome profiling experiments. Coimmunoprecipitation (co-IP) of CFTR bound with interactors was carried out as described previously (25, 40). Briefly, cell lysates were normalized by dilution to a common protein concentration at 1 mL total volume and precleared with 4B Sepharose beads (Sigma Aldrich, St. Louis, MO) at 4 °C for 2 h while rocking. Precleared lysates were then immunoprecipitated with Protein G beads covalently cross-linked to 24-1 antibody (6 mg antibody/mL of beads) overnight at 4 °C while rocking. Beads were washed three times with TNI buffer, twice with TN buffer, all buffer was removed with a needle and then frozen at –80 °C for 1.5 h. Proteins were then eluted off the beads while shaking at 750 RPM at 37 °C for 30 to 60 min with elution buffer (0.2 M glycine and 0.5% IGEPAL CA-630, pH 2.3). The elution step was repeated once and combined to immediately neutralize with a 10:1 ratio of 1 M ammonium bicarbonate solution.
MS sample preparation of co-IP samples was performed as described previously (25, 26). Briefly, samples were chloroform/methanol precipitated, rinsed with methanol, dried, and reconstituted in 1% Rapigest SF (Waters Corporation, Milford, MA). Each sample was reduced with 5 mM TCEP (Sigma Aldrich, St. Louis, MO), alkylated with 10 mM iodoacetamide (Sigma Aldrich, St. Louis, MO), and digested with 0.5 μg of trypsin (Sequencing Grade, Promega, or Pierce) in 50 mM HEPES (pH 8.0) at 37 °C with shaking for a least 10 h. Digested peptides were diluted to 60 μL with water, labeled with 40 μL of TMT pro 16plex reagents (Thermo Fisher) for 1 h, and quenched with 10% w/v ammonium bicarbonate. The 16 TMT-labeled samples were pooled, acidified with MS-grade formic acid (Sigma Aldrich, St. Louis, MO), concentrated using a SpeedVac (Thermo Fisher, Waltham, MA), resuspended in 1,500 μL of buffer A (95% water, 4.9% acetonitrile, and 0.1% formic acid), and loaded onto a triphasic MudPIT column. An Exploris 480 (Thermo Fisher, Waltham, MA) mass spectrometer equipped with an UltiMate3000 RSLCnano System (Thermo Fisher, Waltham, MA) was used for LC-MS/MS analysis as described previously (25, 26).
Proteome Discoverer 2.4 was used for peptide identification and TMT-based quantification as described previously (26). MS/MS spectra were searched using SEQUEST against a UniProt human proteome database (released 03/25/2014) using a decoy database of reversed peptide sequences. The following parameters were employed: 0.02 Da fragment mass tolerance, 10 ppm peptide precursor tolerance, six amino acid minimum peptide length, trypsin cleavage with a max of two missed cleavages, static cysteine modification of 57.0215 Da (carbamidomethylation), and static N-terminal and lysine modifications of 304.2071 Da (TMT pro 16plex). Percolator was used to filter SEQUEST search to minimize the peptide false discovery rate to 1% and require a minimum of two peptides per protein identification. Reporter Ion Quantification processing node in Proteome Discoverer 2.4 was used to quantify TMT reporter ion intensities and a summation was performed for peptides in the same protein.
Luciferase-Based Ribosomal Frameshifting Measurements.
Ribosomal frameshifting measurements were made using a previously described dual-luciferase reporter system that was modified to avoid aberrant splicing artifacts (24). Briefly, all dual-luciferase reporters were generated from a pJD2044 plasmid containing firefly and renilla luciferase and a CMV promoter. Potential cryptic splice sites in the renilla luciferase gene were identified using the human splicing finder webtool and were removed using a series of synonymous codon mutations (246 T>C, 249G>A, 888T>G, and 891G>A) (41). A 138 bp fragment of the CFTR transcript (3,963 to 4,101) was inserted between the 5′ renilla luciferase (0-frame) and 3′ firefly luciferase (−1-frame) genes. P2A linkers were inserted both upstream and downstream of the CFTR insert. HEK293T cells were plated in six-well dishes 1 d prior to transfection of reporter constructs using Lipofectamine 3000 (ThermoFisher Scientific, cat #L3000015). Two days posttransfection, cells expressing dual-luciferase reporter constructs were harvested and lysed using the Passive Lysis Buffer provided in the Dual-Luciferase Reporter Assay System (Promega, cat #E1980). Relative activities of the firefly and renilla luciferase domains in the clarified lysate were then measured using the Dual-Luciferase Reporter Assay System on a Synergy Neo2 plate reader (Biotek, Winooski, VT) in accordance with the manufacturer’s instructions. To determine the –1 ribosomal frameshifting efficiency, the firefly: renilla luciferase ratio was normalized relative to that of a construct lacking the insert as was previously described (42).
CFTR-Mediated hYFP Quenching Measurements.
A fluorescent quenching functional assay based on a previously described method (27, 43) was used to compare the relative functional conductance of CFTR mutants. A pcDNA5 vector bearing an attB recombination site in place of its CMV promoter, untagged CFTR cDNA encoding each variant, and an IRES hYFP-P2A-mKate sensor cassette was used to generate a series of stable HEK293T cell lines that inducibly express each CFTR construct as previously described (38, 39). Recombinant HEK293T cells stably expressing CFTR were grown in six-well dishes and dosed with either DMSO (vehicle) or 3 µM VX-445 and VX-661 48 h after plating. Following 18 h. of treatment with correctors, the cells were harvested with trypsin, washed once in PBS containing 5 mM EDTA (pH 7.4), washed twice in PBS containing 137 mM sodium gluconate (pH 7.4), and then resuspended in PBS containing 137 mM sodium gluconate (pH 7.4), 25 µM Forskolin, 50 µM Genestein, and 10 µM VX-770. Cells were incubated in the resuspension media for 10 min prior to a final wash and resuspension in PBS containing 137 mM sodium gluconate (pH 7.4) prior to analysis. hYFP quenching was initiated with the addition of 25 mM sodium iodide immediately prior to the analysis of single-cell forward light scattering, side light scattering, hYFP fluorescence (488 nm laser, 530/30 nm emission filter), and mKate fluorescence (561 nm laser, 610/20 nm emission filter) intensity values as a function of time using a BD LSRII flow cytometer (BD Biosciences, Franklin Lakes, NJ). Data were analyzed using FlowJo software (Treestar, Ashland, OR). OriginPro 2019 (Origin, Northampton, MA) was used to globally fit the decay of single-cell hYFP: mKate ratios as a function of time using the following exponential decay function:
where y is the hYFP: mKate fluorescence intensity ratio at time t, y0 is the baseline hYFP: mKate fluorescence intensity ratio at t → ∞, A1 is the amplitude of the signal, and k1 is the observed rate constant for the exponential decay. The value of y0 was fixed in the global fit to an experimentally determined value for the hYFP: mKate intensity ratio of cells prior to their treatment with sodium iodide. The quenching half-life was calculated from the globally fit parameters using the following equation:
where t1/2 is the half-life, and k1 is the globally fit rate constant.
Patch-Clamp Measurements.
Patch-clamp recording was used to estimate and compare the channel open probability (Po) of ΔF508 CFTR and ΔF508- RFmut CFTR. Briefly, unitary currents were recorded in excised, inside-out patches from HEK293T cells transiently transfected with either construct. Both ΔF508 and ΔF508- RFmut expressing cells were incubated at 27 °C for 24 to 48 h before patch experiments. Patch pipettes were pulled from Warner Instrument G85150T-3 glass to a tip resistance of 4 to 12 MΩ. Both the pipette and bath solutions were symmetrical and contained 140 mM N-Methyl-D glucamine, 3 mM MgCl2, 1 mM EGTA, and 10 mM TES (pH 7.3). CFTR channels were activated by 1.5 mM Mg-ATP and 123 U/mL of recombinant Protein Kinase A (PKA, Promega, Madison, WI) in the cytosolic bath. Patches were held at +60 mV for unitary current recordings which were analog filtered at 200 Hz and then digitally filtered at 10 Hz with Clampfit 9.2 software (Axon Instruments, San Jose, CA). Data acquisition and analysis were performed using pCLAMP 9.2 software. All patch-clamp experiments were performed at room temperature.
Po values of ΔF508 CFTR and ΔF508- RFmut CFTR were determined from patches as described previously (44, 45). Recordings containing fewer than 8 simultaneous openings prior to the addition of VX-770 (200 nM) were selected for analysis. The number of channels in each patch (N) and the single channel Po for that patch (NPo) were estimated using Clampfit 9.2. The Po value of channels under control conditions was then estimated by dividing the total number of channels (N) in that patch which was estimated from the steady-state macroscopic current measured after potentiator addition; i.e., as N = I/i where I is the steady-state macroscopic current measured after potentiator addition and i is the unitary current at the holding potential of 60 mV (0.4 pA). This provides a minimal estimate of N and a maximal estimate of Po before potentiator addition due to the underlying assumption that the Po for potentiator-activated channels approaches unity.
Detailed methodological information for plasmid preparation and mutagenesis, mass spectrometry data analysis, western blotting, flow cytometry-based immunostaining measurements, and Cas9-mediated gene editing can be found in SI Appendix, Supplemental Methods section.
Data, Materials, and Software Availability
Flow cytometry, western blots, mass spectrometry, luciferase reporter measurements data have been deposited in Mendeley Data (46).
Acknowledgments
We thank Suchetana Mukhopadhyay, Jonathan Dinman, and Jeff Brodsky for helpful critiques of our initial findings. We thank Christiane Hassel for technical input and assistance. We also acknowledge the support of the Indiana University Flow Cytometry Core Facility and the Indiana University Center for Genomics and Bioinformatics. This research was supported in part by grants from the National Institute of General Medical Sciences to J.P.S. (R01GM138845 and R35GM152086). Portions of the paper were developed from the thesis of P.J.C.
Author contributions
P.J.C., F.J.R., A.T., W.W., M.K., E.F.M., A.G.M., C.P.K., S.V.R., L.P., W.D.P., and J.P.S. designed research; P.J.C., F.J.R., A.T., W.W., M.H., M.K., E.F.M., K.N., J.W.-R., J.-L.P., N.B.Z., B.T.P., A.G.M., C.P.K., W.D.P., and J.P.S. performed research; P.J.C., A.G.M., and S.V.R. contributed new reagents/analytic tools; P.J.C., F.J.R., A.T., W.W., M.H., M.K., E.F.M., J.W.-R., J.-L.P., N.B.Z., B.T.P., A.G.M., C.P.K., S.V.R., L.P., W.D.P., and J.P.S. analyzed data; and P.J.C., L.P., W.D.P., and J.P.S. wrote the paper.
Competing interests
The authors declare no competing interest.
Supporting Information
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Copyright © 2024 the Author(s). Published by PNAS. This article is distributed under Creative Commons Attribution-NonCommercial-NoDerivatives License 4.0 (CC BY-NC-ND).
Data, Materials, and Software Availability
Flow cytometry, western blots, mass spectrometry, luciferase reporter measurements data have been deposited in Mendeley Data (46).
Submission history
Received: August 1, 2024
Accepted: August 19, 2024
Published online: October 10, 2024
Published in issue: October 15, 2024
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Acknowledgments
We thank Suchetana Mukhopadhyay, Jonathan Dinman, and Jeff Brodsky for helpful critiques of our initial findings. We thank Christiane Hassel for technical input and assistance. We also acknowledge the support of the Indiana University Flow Cytometry Core Facility and the Indiana University Center for Genomics and Bioinformatics. This research was supported in part by grants from the National Institute of General Medical Sciences to J.P.S. (R01GM138845 and R35GM152086). Portions of the paper were developed from the thesis of P.J.C.
Author contributions
P.J.C., F.J.R., A.T., W.W., M.K., E.F.M., A.G.M., C.P.K., S.V.R., L.P., W.D.P., and J.P.S. designed research; P.J.C., F.J.R., A.T., W.W., M.H., M.K., E.F.M., K.N., J.W.-R., J.-L.P., N.B.Z., B.T.P., A.G.M., C.P.K., W.D.P., and J.P.S. performed research; P.J.C., A.G.M., and S.V.R. contributed new reagents/analytic tools; P.J.C., F.J.R., A.T., W.W., M.H., M.K., E.F.M., J.W.-R., J.-L.P., N.B.Z., B.T.P., A.G.M., C.P.K., S.V.R., L.P., W.D.P., and J.P.S. analyzed data; and P.J.C., L.P., W.D.P., and J.P.S. wrote the paper.
Competing interests
The authors declare no competing interest.
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Ribosomal frameshifting selectively modulates the assembly, function, and pharmacological rescue of a misfolded CFTR variant, Proc. Natl. Acad. Sci. U.S.A.
121 (42) e2414768121,
https://doi.org/10.1073/pnas.2414768121
(2024).
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