Abstract
Proteins synthesized in the cell can begin to fold during translation before the entire polypeptide has been produced, which may be particularly relevant to the folding of multidomain proteins. Here, we study the cotranslational folding of adjacent domains from the cytoskeletal protein α-spectrin using Force Profile Analysis (FPA). Specifically, we investigate how the cotranslational folding behavior of the R15 and R16 domains are affected by their neighboring R14 and R16, and R15 and R17 domains, respectively. Our results show that the domains impact each other’s folding in distinct ways that may be important for the efficient assembly of α-spectrin, and may reduce its dependence on chaperones. Furthermore, we directly relate the experimentally observed yield of full-length protein in the FPA assay to the force exerted by the folding protein in pN. By combining pulse-chase experiments to measure the rate at which the arrested protein is converted into full-length protein with a Bell model of force-induced rupture, we estimate that the R16 domain exerts a maximal force on the nascent chain of ∼15 pN during cotranslational folding.
Significance In living cells, proteins are produced in a sequential way by ribosomes. This vectoral process allows the growing protein chain to start to fold before translation has been completed. Thereby, cotranslational protein folding can be significantly different than the folding of a full-length protein in isolation. Here we show how structurally similar repeat domains, normally produced as parts of a single long polypeptide, affect the cotranslational folding of their neighbors. This provides insight into how the cell may efficiently produce multidomain proteins, paving the way for future studies in vivo or with chaperones. We also provide an estimated magnitude of the mechanical force on the nascent chain generated by cotranslational folding, calculated from biochemical measurements and molecular dynamics simulations.
Introduction
Classically, protein folding has been studied in vitro using purified proteins, providing a detailed picture of how the biophysical properties of a given polypeptide chain affect folding pathways. However, studies of purified proteins do not reflect the vectorial nature of biological protein synthesis, where each amino acid is added sequentially on a time scale that allows the growing protein to explore its continually expanding energy landscape (1).
The discovery (2, 3) and engineering (4, 5) of translational arrest peptides (APs) has provided us with a new tool to examine how proteins fold cotranslationally. To date, the folding of several small proteins and protein domains has been studied using the force-sensitive AP from the E. coli SecM protein, establishing a method we have called Force-Profile Analysis (FPA) (6–15). Recently, we demonstrated that FPA faithfully picks up cotranslational protein folding events observed by direct biophysical measurements (15).
Spectrin, the primary structural component of the cellular cytoskeleton (16–19), is composed of two heterodimers of α- and β-spectrin that come together in a coiled tetramer (20). The 285 kDa non-erythroid αII isoform of spectrin contains 20 three-helix bundle domains, which provide interesting model systems for protein folding. This is because, while they are structurally nearly identical (21), they vary in stability (22–25), folding rate (26, 27), and folding mechanism (28). In particular, folding of the 15th, 16th, and 17th α-spectrin domains (herein called R15, R16, and R17, respectively) has been extensively studied in vitro (24, 27, 29–32).
Previously, using FPA, we demonstrated that R15 and R16 are able to fold cotranslationally and that the point at which this folding begins does not seem to be influenced by the in vitro folding rate (9). Specifically, while R15 folds very quickly in vitro, cotranslational folding of R16 starts at shorter chain lengths than does folding of R15. Indeed, even when the folding core of R15 was substituted into R16, which gives R16 the folding properties of R15 in vitro (33), the protein was still able to fold earlier than R15 during translation. When the minimal, five-residue R15 folding nucleus (34) was substituted into R16, this led to late cotranslational folding, but early folding was restored when this mutant was expressed in tandem with R15, as it would be in vivo. This demonstrated that the mutant R16 variants fold cotranslationally via early intrachain contacts, at chain lengths where the R15 folding nucleus cannot yet form. This was not observed in vitro because the purified mutant R16 protein would preferentially fold using the “fast” R15 nucleus rather than the slower R16 contacts. Cotranslationally, R16 folds using the first available pathway, i.e., the slower R16 folding contacts.
Here, we have extended our studies on spectrin to determine how the presence of neighboring domains affect the folding of R15 and R16. We find that the cotranslational folding of these two domains is differently affected by their respective up- and downstream neighbor domains, a previously unappreciated feature of spectrin folding. We were also able to observe that the synthesis of native, likely helical, structures downstream of the folding spectrin domain increases the observed folding force and further stabilizes the fold even in the presence of amino acid substitutions that destabilize the final native spectrin structure.
While it has been shown previously that the cotranslational folding of proteins can provide a mechanical force to release SecM-mediated ribosomal stalling (6), the magnitude of the force has never been directly measured by FPA. Using pulse-chase experiments, we have now measured the rate at which arrested protein is converted to full-length protein under standard experimental conditions. These rates were fit to a Bell model (35) for force-induced rupture, using parameters elucidated by molecular dynamics simulations (12), to provide an estimate of the magnitude of the pulling force exerted on the nascent chain by cotranslational protein folding. The determined force is in good agreement with previously published measurements using optical tweezers (6).
Results
The force profile analysis (FPA) assay
APs are short stretches of polypeptide that interact with the ribosome exit tunnel in such a way that translation is stalled when the last codon in the portion of the mRNA that codes for the AP is located in the ribosomal A-site (36). APs generally have regulatory functions in cells, controlling, e.g., translation initiation on downstream ORFs in polycistronic mRNAs (36). Interestingly, the stalling efficiency of many APs has been shown to be sensitive to pulling forces exerted on the nascent polypeptide chain, with high pulling force leading to reduced stalling (6, 37, 38). APs can therefore be used as force sensors, reporting on cotranslational events that in one way or another generate pulling forces on the nascent chain (7, 38, 39). The factors influencing such force-generating events were recently examined using molecular simulations and statistical mechanics models, in particular the effect of translation speed (40). Although arrest peptide measurements of force do not include the effect of translation speed directly, this could be accounted for by using a suitable kinetic model for coupled folding and translation (41).
In FPA, a force-generating domain in a polypeptide is placed at increasing distances upstream of an AP, Fig. 1a, and the degree of translational stalling is measured for the corresponding series of protein constructs. Figure 1b shows how FPA can be applied to study the cotranslational folding of soluble protein domains, e.g., tandem spectrin domains. In the construct shown on the left, the chain is long enough for one of the two spectrin domains to have already folded, while the second spectrin domain is largely buried in the exit tunnel and thus cannot fold when the ribosome reaches the C-terminal end of the AP. Therefore, little force is generated on the AP, and stalling is efficient. In the right-hand construct, the chain is so long that both spectrin domains have folded prior to the ribosome reaching the C-terminal end of the AP, and again little force is generated on the AP. In the middle construct, however, the chain is just long enough that the second spectrin domain can begin to fold if the linker that connects it to the AP is stretched beyond its equilibrium length. Under this condition, some of the free energy gained upon folding will be converted to tension in the linker and generate a pulling force on the AP, resulting in reduced stalling and increased production of full-length protein.
The cotranslational folding process can hence be followed by measuring the amount of full-length and arrested protein product at each linker length, Fig. 1c. Folding transitions will appear as peaks in a “force profile” plot of the fraction full-length protein (fFL) vs. linker length (L), Fig. 1d, and can be described by the peak amplitude and the linker lengths that define the onset (Lonset), maximum (Lmax), and end (Lend) of the peak, as shown.
Cotranslational folding of tandem spectrin domains
In the intact spectrin protein, the C-terminal α-helix in one domain is continuous with the N-terminal α-helix in the next domain, Fig. 2a. Consecutive domains are thus intimately connected to each other, and can influence each other’s thermodynamic stability (24, 27, 31, 42). In order to better understand the cotranslational folding of spectrin, we decided to systematically evaluate the effects of up- and downstream domains on the force profiles of the R15 and R16 spectrin domains by analyzing the two- and three-domain combinations shown in Fig. 2b. For each domain combination, a force profile was recorded by generating a series of constructs where the 17-residue SecM(Ec) AP (FSTPVWISQAQGIRAGP) from the E. coli SecM protein was placed at different linker lengths L from the C-terminal end of the R15 or R16 domain, translating each construct for 20 minutes in an in vitro translation system (43) in the continuous presence of [35S]-methionine, separating the translation products by SDS-PAGE, and calculating fFL from the intensities of the bands corresponding to the full-length and arrested forms of the protein. The full amino acid sequences of all constructs are given in Supplementary Table S1.
In order to distinguish which spectrin repeat is being assayed in the tandem constructs, we use the following naming convention. Constructs named “nL” (for LepB linker) are used to follow the folding of most C-terminal spectrin repeat (i.e, R15R16 nL follows the folding of R16) and constructs named “nT” (for Truncated spectrin linker) follow the folding of the penultimate spectrin repeat (i.e. R15R16 nT follows the folding of R15).
The results are shown in Fig. 3a-c. These include previous measurements collected for R16 nL and R15R16 nL (9) along with additional replicates and newly collected data for R15 nL. To facilitate comparison of the results, the Lonset, Lmax, and Lend values extracted from each force profile are summarized in Supplementary Fig. S1; note that Lmax values for force profiles that reach saturation (fFL ≈ 1) cannot be accurately determined. As seen in Fig. 3b, the cotranslational folding of the R15 domain is unaffected by the presence of the upstream R14 domain (dark blue and light blue curves), while the presence of the N-terminal part of the downstream R16 domain induces a reduction in Lonset from 36 to 34 residues and a marked increase in the amplitude of the peak (compare the dark blue and purple curves). Thus, the onset of folding of R15 is sensitive to the presence of the early parts of the N-terminal α-helix of the downstream R16 domain, but is not affected by the presence of the upstream R14 domain.
For the R16 domain, the effects of the neighboring R15 and R17 domains are more dramatic, Fig. 3c. The presence of the upstream R15 domain leads to a reduction in Lonset from 30 to 28 residues (compare the red and magenta curves), as shown before (9). The presence of the downstream R17 domain does not appreciably affect Lonset but leads to an increase in amplitude and a shift to a higher Lend value (compare the red and orange curves). Finally, when flanked by both the R15 and R17 domains, the R16 folding transition starts at a lower Lonset and ends at a higher Lend than for the isolated R16 domain (compare the red and brown curves).
In order to more precisely define Lmax for constructs were fFL approaches 1, we substituted two amino acids in the relatively weak SecM(Ec) AP to create the stronger SecM(Ec-sup1) variant (FSTPVWISQAPPIRAGP) (44) in the R16 nL, R15R16 nL, R16R17 nT, and R15R16R17 nT constructs, Fig. 3d. Again, the presence of R15 reduces Lonset and Lmax by ∼2 residues (note that the R15R16 nL and the R15R16R17 nT profiles overlap perfectly in the interval L = 23-31), and the presence of R17 increases Lend by 6-7 residues (see Supplementary Fig. S2). Notably, the bi-modal shape of the R15R16R17 nT profile with peaks at L = 29 and 37 residues is an almost perfect match to the sum of the R15R16 nL and R16R17 nT profiles, Supplementary Fig. S2f, suggesting that the R16 part folds first (stabilized by the R15 C helix, c.f., Fig. 2a), followed by a second folding event involving the R17 A helix (stabilized by the R16 C helix).
Interestingly, both R16 folding constructs that include parts of R17 (R15R16R17 nT and R16R17 nT) show an increase in fFL at the longest linker length (L = 61 residues), Fig. 3c. At this point the entire R17 A helix, the loop, and the very beginning of R17 helix B have been translated (see Table S1). This increase in fFL may thus signal an early interaction between helices A and B in R17.
As a control, we introduced mutations that are known to inhibit folding of R15 and R16 in vitro (45, 46) into the R15 nL, R15R16 nT, R16 nL, R16R17 nT, and R15R16R17 nT constructs, Fig. 4 and Supplementary Fig. S3. As expected, fFL values for the “non-folding” R15(nf) nL and R16(nf) nL constructs were strongly reduced, but, surprisingly, fFL values remained high for linker lengths between Lmax and Lend for R15(nf)R16 nT, R16(nf)R17 nT, and R15R16(nf)R17 nT constructs. Since the difference between the nL and nT series of constructs is the presence of a C-terminal “linker” derived from the N-terminal part of R16 or R17 respectively (replacing unstructured segments from LepB), this suggested that the linker itself, perhaps together with C-terminal parts of the upstream spectrin domain, might fold in the nf-mutants.
To investigate this possibility, we started from the R15R16 nT, L = 40 construct (whose corresponding nf-mutant has a much higher fFL value than the R15(nf) nL version, Fig. 4a) and successively removed helices A, B, and C from the R15 part (i.e. N-terminal truncations), Fig. 4b. We found that removing helix R15-A or both helices R15-A and R15-B decreased fFL, but only to 0.47 and 0.36, respectively, Fig. 4c (compare points #4, to #3, and #2). Only with the removal of all of R15, leaving only 19 amino acids of the R16 A helix, did fFL decrease to baseline (point #1). Thus, the cotranslational formation of a continuous helical structure encompassing parts of the R15 C and R16 A helices appears to cause an increase in fFL at L = 40; indeed, it is known from previous work that helices can form cotranslationally in the exit tunnel (47). The introduction of helix-breaking residues into the middle of the R16 A helix (see Table S1) in the R15-ABC/R16-A (point #4) and R15-BC/R16-A (point #3) constructs decreased the fFL to the same level as R15-C/R16-A (point #2), Fig. 4c (open circles). Likewise, comparison of constructs where the R16 A helix in the R15R16 nT and R15(nf)R16 nT, at L = 40 was replaced by a non-helical segment from LepB at a comparable length (L = 39 and L = 41, respectively) led to reductions in fFL (compare points #5 & 6 and #4 & 7). Similar behavior was also observed for the R16R17 nT, L = 43 constructs, Supplementary Fig. S3, implying that the R17 A helix forms a cotranslational folding intermediate together with the R16 C helix, as suggested above.
We conclude that the cotranslational folding of the R15 domain is affected by its downstream but not by its upstream neighbor domain, and that folding of R16 is affected both by its upstream and downstream neighbors. Different spectrin domains thus not only fold via different folding mechanisms, but their cotranslational folding is also differently impacted by their up- and downstream neighbor domains, a previously unappreciated feature of spectrin folding. We attribute the contribution of the downstream neighbor to the formation of helical structure encompassing helix C from the upstream domain and helix A from the downstream domain.
A quantitative relation between fFL and pulling force
In all FPA studies published to date, the quantitative relation between the calculated fFL values and the underlying pulling force acting on the nascent chain has remained undefined (although attempts have been made to derive it from simulations or other kinds of theoretical modeling (11, 39)). Using the PURE translation system, Goldman et al. (6) showed that the interactions between the SecM(Ec) AP and the ribosome exit tunnel can be disrupted by a mechanical force applied through optical tweezers, and that the rate by which the translational stall induced by the SecM(Ec) AP is released, kR, increases in step with the amount of force, F, applied to the nascent chain, in a way that can be approximated by the Bell model (35) for force-induced rupture: where k0 = 3 × 10−4 s−1 (95% confidence interval (CI): 0.5 × 10−4 s−1, 20 × 10−4 s−1) is the release rate at zero pulling force, and Δx‡ = 0.4 nm (95% CI: 0.1 nm, 0.8 nm) is the distance to the transition state.
The work by Goldman et al. (6) suggested to us that approximating kR with the rate of conversion of the arrested form of a given construct to the full-length form as measured in a pulse-chase experiment would allow us to estimate the corresponding pulling force using the relation in Eq. [1] between F and kR. To this end, we did pulse-chase experiments on a range of spectrin R16 and ADR1a (7) constructs that have fFL values between 0.2 and 0.9, and fit the pulse-chase data to a first-order kinetic equation, Supplementary Fig. S4.
To derive an analytical relation between fFL and F, we reasoned that if fFL were measured at a single delay time (Δt) in a pulse-chase experiment, rather than under the standard continuous-labeling experimental conditions, the release rate (and hence F, according to Eq. [1]) could be estimated from:
Since in the standard experiment translation can be initiated at any time during the 20 min. labelling period, the delay time Δt varies from one ribosome-nascent chain complex to another. Although many factors could in principle contribute to the distribution of delay times, it turns out empirically that using an average delay time Δt = 550 s, approximately equal to half of the total incubation time, describes remarkably well the relation between the standard fFL values (20 min. continuous [35S]-Met labeling) and the release rates measured by pulse-chase experiments, Supplementary Fig. S5.
Combining Eqs. [1] and [2], one can solve for the pulling force F:
To obtain good estimates of the Bell parameters k0, Δx‡ we started from the values obtained in (6) and performed a local optimization by requiring that k0 (the release rate at zero force) is close to kR measured for the “zero-force” construct R16 nL27, Fig. 5a, and that the titin I27, spectrin R16, and ADR1a force profiles in (12) are well reproduced by the molecular dynamics simulation also described in (12). These conditions are both fulfilled by setting k0 = 3 × 10−4 s−1 (i.e., the same value as in (6) and equal to kR for R16 nL27, Fig. 5a) and x‡ = 0.65 nm (well inside the 95% CI from (6)), Supplementary Fig. S6. With these parameter values and Δt = 550 s, Eq. [3] nicely captures the relation between the force estimated from the measured kR and standard fFL values, Fig. 5b, and can hence be used to predict F from fFL.
Over the interval 0.2 < fFL < 0.9, the experimental data is also well approximated by the simple linear fit F = 22 fFL – 2.2 (blue line). Obviously, Eq. [3] holds only for the 17-residue SecM(Ec) AP used here, and not for other APs of different stalling strengths (39). Presumably, Eq. [3] may be applied also in other contexts where the SecM AP is used to measure pulling forces, such as during membrane protein synthesis or translocation of charged residues across energized membranes (38, 39), although the parameters would first need to be verified to hold for in vivo experiments. We note that another approach to determining Bell parameters under a given set of conditions would be to repeat the FPA experiments using multiple APs with different resistance to force. A global fit of fFL obtained for each AP as a function of linker length L would allow simultaneous determination of the forces exerted by the protein (independent of AP) as well as AP-dependent Bell parameters. In Supplementary Fig. S7 we illustrate this approach using data for translocon-mediated transmembrane helix insertion into the inner membrane of E. coli from in vivo AP measurements.
The force F estimated from pulse-chase measurements or fFL values represents the constant force that would have the same effect on the escape rate from arrest as the combined effect of the forces from both folded and unfolded states. We can compare this inferred force with the ensemble average force calculated from MD simulations, ⟨FMD⟩ = pUFU + pFFF, where pU(F) and FU(F) are, respectively, the population of, and force exerted by, the unfolded (folded) state in the simulation. In previous work (12), we calculated both fFL as well as ⟨FMD⟩ directly from MD simulations. In Supplementary Fig. S8a, we show the relation between ⟨FMD⟩ and fFL from our previous simulations (12), which is in good agreement with the relation between F and fFL calculated from Eq. [3]. This agreement also confirms that the pulling forces are small enough that the force F calculated from simulated fFL values using Eq. [3] is close to the ensemble-average force ⟨FMD⟩ determined by MD simulations; the two forces are not equal in general because of the non-linear relation between release rate and force, such that states exerting a larger force (e.g. the folded state) contribute disproportionately to the average release rate and hence to F (a direct comparison between ⟨FMD⟩ from the MD simulations and F calculated from Eq. [3] is given in Supplementary Fig. S8b).
We note that the forces exerted by a protein folding as it exits the ribosome are qualitatively different from the tensile forces such as those exerted when proteins fold in AFM experiments (as have been performed on spectrin before (48)). Therefore, the force magnitudes probed by the two experiments cannot be directly compared, despite their apparent resemblance. For example, even a small force exerted on the termini of an unfolded protein by an AFM can massively slow the refolding rate because of the large distance the protein must contract against this force in order to fold. The same force magnitude would have a much smaller effect on the refolding rate of a protein attached at one end to the ribosome (11).
Discussion
α-spectrin contains over 20 repeat domains and is produced as a single, long polypeptide. Previously, we observed that the R16 domain starts to fold while a part of its C-terminal α-helix is still in the ribosome exit tunnel (9). We now find that the cotranslational folding of R16 is affected both by the presence of its upstream R15 domain and when the N-terminal part of the following R17 domain is present in the exit tunnel. In contrast, folding of the R15 domain is not affected by the upstream R14 domain, but starts at shorter linker lengths when the N-terminal end of the following R16 domain is present.
In vitro, R16 is stabilized by ∼1.7 kcal/mol by the presence of R15 (49), meaning that, during cotranslational folding, R16 can “sacrifice” a few interactions in the folding nucleus and start to fold at a shorter linker length in the R15R16 nL constructs (9).
For R15, the situation is a bit different because, in contrast to R16, its folding nucleus involves residues located at the C-terminal end of helix C (45). The presence of the upstream R14 domain thus may have little impact on Lonset, because the whole R15 domain anyway must have emerged from the exit tunnel before folding can start. On the other hand, the R15 C helix is stabilized by the presence of the early parts of the R16 A helix, possibly allowing the folding nucleus to form at a shorter linker length and reducing Lonset in the R15R16 nT constructs.
The picture that emerges is that the spectrin domains fold one after the other as they emerge from the exit tunnel, but not completely independently of one another. As seen for the R16 domain, the presence of an already folded N-terminal upstream neighbor domain not only can increase the thermodynamic stability of the folded state of an emerging domain, but can also allow the emerging domain to start folding while still partly buried in the exit tunnel.
Likewise, when the C-terminal α-helix in the emerging domain is extended by a part of the N-terminal α-helix in the following domain, the folding transition can persist to longer linker lengths (as seen for R16) or start at shorter linker lengths (as seen for R15). Finally, the increase in fFL seen for the R16R17 nT and R15R16R17 nT constructs at L = 61 residues (Fig. 3c) is suggestive of an early folding intermediate in R17. Further work is required to investigate this, but it may be possible that the regular helical structure of the spectrin repeats drives a nearly-continuous folding reaction, punctuated by periods when the nascent chain is lengthened but no residues are added to the folded part emerging from the ribosome. In this scenario, at most a short stretch of unfolded polypeptide would be exposed outside the ribosome at any one time, which could minimize the need for protection of the nascent chain by chaperones.
Finally, a pulse-chase analysis has allowed us to measure the release rate kR from the stalled state for different R16 and ADR1a constructs, making it possible to estimate the magnitude of the pulling forces exerted on the AP for different linker lengths, Fig. 5a. This in turn makes it possible to derive expressions for how the magnitude pulling force F depends on fFL (as obtained from our standard continuous-labeling experimental protocol), Eqs. [2] and [3]. In general, we find that the folding of protein domains such as spectrin R16 and ADR1a can generate a maximal force of 15-20 pN on the AP, Fig. 5, in line with theoretical estimates based on MD simulations (40, 50).
Materials and Methods
All the enzymes used for molecular biology were obtained from New England Biolabs. The PUREfrex in vitro transcription-translation system was produced by Eurogentec and purchased via BioNordika. The PURExpress in vitro transcription-translation system was purchased from NEB. GeneJet PCR clean-up kit was purchased from Thermo-Fisher Scientific. Precast NuPAGE gels and running buffers were purchased from Invitrogen. Homemade gels were cast using acrylamide-bisacrylamide mix, Tris, and glycine from VWR. Filter paper for gel drying was from Whatman. All other chemicals were purchased from Sigma.
Cloning
The cDNAs for spectrin R15 and R16 domains were kindly provided by Dr. Jane Clarke and the spectrin R17 and R14 domains were ordered as DNA fragments from Eurofins Genomics GmbH. All protein sequences used in this publication are presented in Supplementary Table S1. We used two different construct types: those with an unrelated linker sequence, labelled nL, and those with a truncated spectrin linker sequence, labelled nT (Fig. 2b). For nL constructs, the respective spectrin domain or domains were cloned into the pET19b plasmid upstream from a Ser-Gly-Ser-Gly sequence attached to a linker sequence derived from the P2 domain of leader peptidase (LepB) followed by the 17-residue AP from the E. coli SecM protein (FSTPVWISQAQGIRAGP) herein referred to as SecM(Ec), and a further 23 amino acids derived from LepB. The shortest constructs (nL/nT 21) include only the SGSG sequence fused to the SecM(Ec) AP. For nT constructs, the respective spectrin domains were cloned into the same pET19b plasmid without a LepB linker sequence, but including the SGSG sequence immediately before the SecM(Ec) AP. The most C-terminal spectrin domain was then sequentially truncated via partially overlapping inverse PCR. For some constructs, a full-length (FL) control was created by changing the critical Pro at the C-terminal end of the SecM AP to Ala, thereby abolishing arrest (51) and yielding only full-length protein. The SecM(Ec-sup1) variants were created by site-directed mutagenesis of the constructs described above. N-terminal truncations of R15 and R16 were carried out by partially overlapping inverse PCR and insertion of the helix breaking insertions was carried out by site-directed mutagenesis where the inserted residues were included in the primer sequence. All sequences were confirmed by DNA sequencing (Eurofins Genomics GmbH).
In vitro expression for Force Profile Analysis (FPA)
Expression and analysis was carried out as described previously (9). Briefly, a linear DNA product is created from each construct plasmid by PCR using Q5 polymerase with forward and reverse primers that anneal to the T7 promoter and terminator regions, respectively. Following PCR clean-up (using the manufacturer’s instructions) the product is confirmed by agarose gel electrophoresis. In vitro transcription and translation is carried out in either the PUREfrex of PURExpress commercial systems (mixed according to the manufacturer’s recommendations). 1 µL of the PCR product and ∼10 µCi [35S]-methionine are mixed for a 10 µL PUREfrex reaction or 0.5 µL of PCR product and ∼5 µCi [35S]-methionine are mixed for a 5 µL PURExpress reaction, followed by incubation at 37°C for 20 min at 600 rpm shaking. Translation is halted by the addition of an equal volume of ice-cold 10% TCA, followed by incubation on ice for at least 30 min. Total protein is sedimented by centrifugation at 4°C for 5 min at 20,000xg. The supernatant is carefully removed and the pellet is resuspended in a suitable volume of 1X SDS-PAGE sample buffer (62.5 mM Tris-HCl pH 6.8, 10% glycerol, 2.5% SDS, 5% β-ME, 0.02% bromophenol blue, and 25 mM NaOH (NaOH is added to neutralize any remaining TCA) by shaking at 37°C and 1,000 rpm for at least 5 min. The prolyl-tRNA that remains attached due to SecM arrest is digested by the addition of 4 µg of RNAse I (2 µL of a 4 µg/µL solution), followed by incubation at 37°C and 600 rpm for 30 min.
Quantifications
Following a brief centrifugation to remove any remaining insoluble material, the sample is loaded onto an appropriate SDS-PAGE gel (12% tris-glycine gels were used for two- and three-spectrin domain constructs and 16 or 18% tris-tricine gels were used for single-spectrin domains and the N-terminal truncations). Following electrophoresis, the gels are dried onto thick filter paper by heating under vacuum (BioRad Model 583 or Hoefer GD 2000), a radioactive molecular weight ladder included in the gel is visualized by spotting the filter paper with a ∼1:1,000 solution of [35S]-methionine in 1X SDS-PAGE sample buffer, and the gel is exposed to a phosphorimager screen for 12 to 72 hours depending on the strength of the signal. The screen is imaged using a Fujifilm FLA9000 (50 µm pixels), and densitometry analysis on the resultant raw image (TIFF format) file is carried out using FIJI (ImageJ) software. The densitometry values are quantified using our in-house EasyQuant software and the fraction full-length protein, , is calculated from the intensities of the full-length (IFL) and arrested (IA) bands. See Supplementary Fig. S9 for examples of gels. Independent replicate in vitro translation reactions were carried out for all spectrin constructs in the folding peaks, see Supplementary Table S2 (191 unique constructs and 405 independent data points). The majority of the data points in the folding peaks were collected in triplicate, with the remaining points collected in duplicate. A small number of data points outside of the folding peaks were collected as single measurements to save costs.
Pulse-chase experiments
The rate at which translation recommences following the arrest of various constructs was measured for five R16 constructs and two ADR1 constructs (7) chosen to represent the range of fFL values measured during typical experimental conditions. Pulse-chase experiments were carried out using the PURExpress system. 75 µL reactions were mixed according to the manufacturer’s instructions, with the addition of [35S]-methionine. Following a 5 min incubation at 37°C (pulse) an excess of unlabeled methionine was added and incubation at 37°C was continued. At discrete time points, a 10 µL aliquot of the reaction was removed and mixed with 15 µL of ice-cold 10% TCA and further processed as above. Three independent replicates were collected for each construct (see Supplementary Table S2). The rate calculated is for the conversion of arrested protein to full-length protein:
The rate of this irreversible conversion can be calculated using a 1st order equation:
Where fA(t) is the fraction arrested protein at a given time, t, and fA(0) is the fraction arrested protein at t = 0. Prism 8 (GraphPad Software) was used to calculate the non-linear regression fit of the kinetic equation to each data set (R16 nL-27, -29, -31, -33 and -37, and ADR1 L25 and 27 (7)). The rates were used to calculate the force exerted by the folding domain on the arrested nascent chain as detailed in the main text.
Optimization of the Bell parameters
The coarse-grained molecular dynamics (MD) simulations of cotranslational folding of R16 and ADR1a constructs in WT, ΔuL23, and ΔuL24 ribosomes used here were originally reported in (12). For each construct and each linker length, these simulations were used to calculate the ensemble average force ⟨FMD⟩ = pUFU + pFFF, where pU(F) and FU(F) are, respectively, the population of, and force exerted by, the unfolded (folded) state in the simulation. In order to determine the Bell parameters k0 and Δx‡ in Eq. [1] in the main text, we set k0 = 3 × 10−4 s−1 (i.e., the same value as estimated in (6), and equal to kR for the “zero force” construct R16 nL27, Fig. 5a) and explored a range of Δx‡ values around the approximate value Δx‡ = 0.4 nm given in (6). For each Δx‡ value, we calculated an average kR over unfolded and folded states as kR = pUk0 exp[FU Δx‡/kBT] + pFk0 exp[FF Δx‡/ kBT] and then obtained fFL (setting Δt = 550 s) from Eq. [2] in the main text. The simulated fFL values (i.e, the simulated force profiles) obtained in this way were then compared with the experimental force profiles reported in (12) for the I27 and R16 domains expressed with WT, ΔuL23, and ΔuL24 ribosomes, and for the ADR1a domain expressed with WT and ΔuL24 ribosomes. The parameter values k0 = 3 × 10−4 s−1, Δx‡ = 0.65 nm were found to give the best fit between the simulated and experimental force profiles (see Supplementary Fig. S7).
Author contributions
GK, OBN, and GvH conceived the project. GK and OBN designed and performed the experiments. PT and RB performed the theoretical calculations. All authors contributed to writing the manuscript.
Competing Interests
The authors declare no competing interests.
Data Availability
The quantified raw data points are provided in Supplementary Table 2.
Acknowledgements
This work was supported by grants from the Knut and Alice Wallenberg Foundation (2012.0282), the Swedish Cancer Society (15 0888), and the Swedish Research Council (621-2014-3713) to GvH, and from the Intramural Research program of the National Institute of Diabetes and Digestive and Kidney Diseases of the NIH to RBB, and from NIH (grant R35GM122543) to FPA. This work utilized the computational resources of the NIH HPC Biowulf cluster. (http://hpc.nih.gov). We thank Dr. Jane Clarke and Dr. Adrian Nickson for providing plasmids with spectrin DNA and Dr. Rickard Hedman for programming and maintenance of the EasyQuant software.
Footnotes
Abbreviations: PCR, polymerase chain reaction; TCA, trichloroacetic acid; SDS-PAGE, sodium dodecylsulphate polyacrylamide gel electrophoresis; PTC, peptidyl transferase center
New data in Fig. 3 and Fig. S2