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Structural Biology Unit, Center for Cooperative Research in Biosciences (CIC bioGUNE), Basque Research and Technology Alliance (BRTA), Bizkaia Technology Park, Derio, SpainDepartamento de Bioquímica and Instituto Biofisika, Consejo Superior de Investigaciones Científicas–Universidad del País Vasco/Euskal Herriko Unibertsitatea (CSIC, UPV/EHU), Bizkaia, SpainIKERBASQUE, Basque Foundation for Science, Bilbao, Spain
Fold-switch pathways remodel the secondary structure topology of proteins in response to the cellular environment. It is a major challenge to understand the dynamics of these folding processes. Here, we conducted an in-depth analysis of the α-helix–to–β-strand and β-strand–to–α-helix transitions and domain motions displayed by the essential mannosyltransferase PimA from mycobacteria. Using 19F NMR, we identified four functionally relevant states of PimA that coexist in dynamic equilibria on millisecond-to-second timescales in solution. We discovered that fold-switching is a slow process, on the order of seconds, whereas domain motions occur simultaneously but are substantially faster, on the order of milliseconds. Strikingly, the addition of substrate accelerated the fold-switching dynamics of PimA. We propose a model in which the fold-switching dynamics constitute a mechanism for PimA activation.
Proteins do not just occupy a single state but are in dynamic exchange between a set of conformations, and some of these states may at times be populated at very low levels, so-called invisible or dark states (
). Motions occur on a broad range of timescales, and growing evidence suggests that dynamics are instrumental for the function of a multitude of proteins. In enzymatically mediated reactions, protein dynamics enable the recruitment and binding of substrates, stabilization of transition states, and adaptation to a dynamic membrane environment or binding partners (
). Fold-switching, i.e. the interconversion of secondary structure elements, is a functionally essential dynamic event for a certain class of proteins. Such motions modify the topology of a protein, including α-helix–to–β-strand and/or β-strand–to–α-helix transitions (
). Alterations of such dynamic processes are essential for and causative of human disease, as in the case of the prion protein PrP and a variety of proteins associated with neurodegenerative diseases, including α-synuclein, amyloid β-peptide, and huntingtin (
). PimA catalyzes the first step in the biosynthesis of phosphatidyl-myo-inositol mannosides (PIMs) by adding a mannose residue donated by the nucleotide sugar GDP-mannose (GDP-Man) to a phosphatidyl-myo-inositol lipid, which is anchored into the cytoplasmic phase of the plasma membrane (
). In the crystal structure of unliganded PimA, two distinct conformations of a region in the N-terminal domain, termed the reshuffling region (residues 118–163), are seen, suggesting that this region can undergo motions between a compact and an extended conformation, even in the absence of substrate (Fig. 1A) (
). Moreover, the crystal structures of PimA in the presence of GDP or GDP-Man support the hypothesis that the reshuffling region displays fold-switching of secondary structure elements with both α-helix–to–β-strand and β-strand–to–α-helix transitions upon addition of substrate (Fig. 1A) (
). Finally, the occurrence of an open-to-closed motion between the N- and C-terminal Rossmann fold domains has been predicted and experimentally demonstrated to occur in PimA and other GT-B glycosyltransferases (
Here, we demonstrate by using 19F NMR methods that motions of PimA occur on a second-to-millisecond timescale and include interconversions between several conformational states. Our observations provide a basis to suggest an activation mechanism for PimA in which the addition of substrate accelerates dynamics between active and inactive conformations.
19F labeling is a sensitive approach to study dynamics of PimA by NMR
19F nuclei are NMR active with a gyromagnetic ratio that is 0.94 times that of protons, have 100% natural abundance, and display high sensitivity to their magnetic environment (
). At the same time, fluorine is nearly absent from biological molecules, reducing the number of resonances observed in a spectrum to those that stem from artificially introduced 19F labels. Therefore, 19F nuclei are suitable probes that allow employing NMR experiments for dynamic studies even for large proteins for which more common labeling approaches are not successful (
). Based on analyses of the different crystallographic structures of PimA (Fig. 1A), we introduced a Cys mutation in position Arg144 (here termed PimAR144C) (Fig. 1A) and labeled it with 1,1,1-trifluoroacetone (TFA), giving rise to a 19F labeled mutant PimAR144C-TFA (see Experimental procedures). The introduced 19F label of the mutant PimAR144C-TFA is located in different chemical environments in the different crystallographic conformations of PimA, as highlighted in Fig. 1A. In the active state, the residue is buried inside the protein and, thus, not exposed to solvent, while it is solvent-exposed in the inactive states. 19F nuclei have been shown to be sensitive to such differences in the hydrophobicity of their immediate environment (
). Thus, it can be expected that the 19F probe located at Cys144 can sense motions that we hypothesize to occur in the reshuffling region, which are 1) extended to compact domain motion and 2) fold-switching dynamics between the inactive and active states.
Given that the probe experiences sufficiently distinct chemical environments, domain motions that are typically on the order of milliseconds can be probed by NMR relaxation dispersion experiments (
We first determined that the introduced label does not alter the three-dimensional structure of PimA. Supporting this notion, wild-type PimA, PimAR144C, and labeled PimAR144C-TFA mutants displayed similar far-UV CD spectra (Fig. S1A) (
). The midpoint temperature of unfolding for the unliganded protein is referred to as Tm. It is expected that the addition of ligands to the protein sample will increase the Tm. The higher the affinity of a ligand, the higher the apparent Tm value, because it requires more energy to dissociate higher affinity compounds (
). As depicted in Fig. S1B, thermal unfolding followed by the far-UV CD signal at 222 nm indicated slight differences in protein stability between unliganded WT PimA, PimAR144C, and PimAR144C-TFA. Interestingly, after the addition of GDP, wild-type PimA, PimAR144C, and PimAR144C-TFA displayed a clear increase in the Tm values (Table S1), supporting the notion that the mutation and labeling do not significantly perturb the enzymatic structure and function (Fig. S1).
Active and inactive states of PimA show distinct resonances
PimA contains two Cys residues buried inside the protein core that are not expected to be labeled during the labeling procedure (see Experimental procedures). To verify this, the labeling scheme was applied to WT PimA. Only resonances that are less than 5 Hz wide were observed in a 19F spectrum (Fig. S2), showing that they stem from small molecules. The resonances could be assigned to residual 3-bromo-1,1,1-trifluoroacetone (BTFA) label and 1,1,1-trifluoroacetone (TFA), the latter being the product of an incomplete labeling reaction. The resonance at −119.8 ppm stems from an impurity in the BTFA solution. Therefore, no labeling was observed for WT PimA. We conclude that PimAR144C-TFA is only labeled at the introduced Cys residue.
Despite being singly labeled, two resonances are observed at −84.1 ppm and −82.7 ppm (Fig. 1B, bottom). As discussed below, these two peaks display saturation transfer, further corroborating that they stem from the same 19F label. To assign the resonances to conformations of PimA, we investigated the 19F spectrum of the triple mutant PimAT126C-V359C-R144C-TFA, previously shown to be locked in the inactive conformation. This variant cannot undergo reshuffling due to a disulfide bond formed between Cys126 and Cys359; these residues are in close spatial proximity in the inactive state but far apart in the active state (
). PimAT126C-V359C-R144C-TFA displays only a single resonance at −84.1 ppm (Fig. 1B, top), which therefore can be assigned to the inactive state. Consequently, the resonance at −82.7 ppm is assigned to the active state. This assignment is corroborated by the fact that the resonance at −84.1 ppm shifts 0.1 ppm upfield in 100% D2O, whereas the resonance at −82.7 ppm does not shift (Fig. 1B, center). For fully solvent-exposed 19F labels, upfield solvent isotope shifts of 0.2–0.3 ppm have been observed when comparing shifts in a 90% H2O/10% D2O to 100% D2O solvent (
), indicating that the resonance at −84.1 ppm stems from partially exposed 19F label. In the crystal structure, Arg144 is shielded from solvent in the active state, whereas this is not the case for the inactive conformations (Fig. 1A), supporting the assignment. The assignment is further supported by extensive molecular dynamics (MD) simulations performed on the different states of PimA labeled with TFA. Based on these simulations, we calculated the solvent-accessible surface area (SASA) of the TFA label in the different states. A higher SASA value indicates that the label is more solvent-exposed. As shown in Fig. S3, the SASA of the TFA label in the active state is 19.3 Å2, which differs markedly from the values obtained for the protein in the inactive forms (96.7 Å2 and 60.1 Å2 for the extended and compact structures, respectively). Thus, MD simulations revealed that the TFA label in the active state is less surface exposed than that in the inactive state, in agreement with the NMR experiments.
PimA undergoes slow conformational exchange between the inactive and active states
To test whether the two resonances in the 19F spectrum of PimAR144C-TFA are in slow exchange, we conducted saturation transfer experiments at increasing saturation times. Exchange rates were obtained from a fit to the model given in (Eq. 4), (Eq. 5), (Eq. 6), (Eq. 7). Experimental data to obtain R1 longitudinal relaxation rates that are required for the analysis are shown in Fig. S4 and Table S2. Figure 2 shows resonance intensity attenuation due to slow conformational exchange for the inactive and active state resonances upon saturation of the active or inactive state resonance, respectively.
In the absence of GDP-Man, the exchange rate was determined to be 2.3 s−1. These dynamics correspond to fold-switching between the active and inactive states of the protein, as demonstrated by the assignments (Fig. 1B). The second timescale of the motion is in accordance with studies of the chemokine lymphotactin, for which fold-switching dynamics were shown to be on the order of seconds (
I, the inactive state; A, the active state, i.e. kIA is the exchange rate from the inactive to the active state, and ΔGIA is the free energy difference between the inactive and active state and vice versa.
ΔGIA (kcal mol−1)
1.08 ± 0.05
0.47 ± 0.04
2.31 ± 0.09
−0.070 ± 0.090
4.2 ± 0.2
0.34 ± 0.04
13 ± 1
−0.390 ± 0.180
1.23 ± 0.04
0.53 ± 0.04
2.31 ± 0.09
0.070 ± 0.090
8.3 ± 0.9
0.66 ± 0.12
13 ± 1
0.390 ± 0.180
a I, the inactive state; A, the active state, i.e. kIA is the exchange rate from the inactive to the active state, and ΔGIA is the free energy difference between the inactive and active state and vice versa.
To test how GDP-Man affects these dynamics, we first conducted a titration experiment to corroborate that the sugar donor saturates the binding site. As shown in Fig. 1C, titration of GDP-Man to 100 μm PimAR144C-TFA leads to a gradual decrease in intensities of both resonances up to a nucleotide sugar concentration of 200 μm. At this concentration, the inactive state resonance shifts 0.2 ppm upfield, and further addition of GDP-Man does not alter the spectrum anymore. At the field strength employed in Fig. 1C (16.4 T), the active state resonance is too broad to clearly observe a resonance shift. Since the line broadening due to the field-dependent chemical shift anisotropy of 19F is reduced at lower field strength, we compared the spectra of PimAR144C-TFA in the presence and absence of GDP-Man at 14.1 T. At this lower field, a downfield shift of the active state resonance of 0.1 ppm was observable at 500 μm GDP-Man and 100 μm PimAR144C-TFA (Fig. S5). Moreover, both peaks broaden upon addition of GDP-Man, indicating that the presence of GDP-Man alters dynamics and/or populations of PimA. This process may be related to GDP-Man binding itself or to the dynamic processes observed in the reshuffling region of PimA, as discussed below. GDP-Man and GDP were previously seen to bind PimA in enthalpy-driven reactions with dissociation constants (KD) of 0.23 μm (ΔH = –15.6 kcal·mol−1) and 0.03 μm (ΔH= –14.0 kcal·mol−1), respectively (
). Thus, the submicromolar KD indicates that under the experimental conditions employed here, 500 μm GDP-Man and 100 μm PimAR144C-TFA binding of GDP-Man to PimA is fully saturated. Thus, we can exclude any contribution of GDP-Man binding to the observed dynamics discussed subsequently.
Under the condition that GDP-Man is present at saturating concentrations, the fold-switching dynamics of PimAR144C-TFA are strongly enhanced, and the exchange rate was observed to increase to 13 s−1 (Fig. 2 and Table 1). A concurrent shift in the population equilibrium toward the inactive states was observed (by around 10%), as evidenced by analyzing peak integrals as well as the saturation transfer experiments (Fig. 2, Fig. S5, and Table S3).
The inactive state of PimA undergoes millisecond motions between a compact and an extended state
To capture possible dynamics on the millisecond timescale, we conducted Carr-Purcell-Meiboom-Gill (CPMG) relaxation dispersion experiments on PimAR144C-TFA at two magnetic field strengths in the absence and presence of saturating amounts of GDP-Man. Assuming a two-state exchange, we fitted the data to Equation 3 (see Experimental procedures) to obtain an estimate of the size of the populations, the exchange rate between them, and the chemical shift difference due to the conformational exchange. Data that were acquired at one magnetic field strength only were fitted to Equation 2 to obtain an apparent exchange rate.
The apparent exchange rate of the resonance representing the inactive state of PimAR144C-TFA and the triple mutant PimAT126C-V359C-R144C-TFA locked into the inactive state are identical (Fig. S6). Although the triple mutant cannot undergo reshuffling to the active state, the faster domain dynamics of the inactive state itself are not affected. This is expected from inspecting the structure of PimA. While the disulfide bond between Cys126 and Cys359 prevents secondary structure reshuffling, it does not impede the extended-to-compact motion of the inactive states (Fig. 1A). Thus, the relaxation dispersion displayed by the inactive state resonance monitors the extended-to-compact dynamics of the reshuffling region.
As shown in Fig. 3A, the resonance in the 19F spectrum of PimAR144C-TFA monitoring the inactive state displays relaxation dispersion between two conformations with different populations, and the profile is affected by the addition of saturating amounts of GDP-Man, reflecting changes in dynamics and population equilibria. In the absence of GDP-Man, the exchange rate is 8000 s−1, which decreases to 5100 s−1 in the presence of GDP-Man (Table 2). Similar exchange rates have previously been observed for motions of a helix of the β2-adrenergic receptor (
). In the absence of GDP-Man, one of the states is highly populated and the other one is not (1% versus 99%). Upon addition of GDP-Man, the less populated of the inactive states becomes slightly more populated (3% versus 97%), which is accompanied by an upfield resonance shift of the inactive state resonance (Fig. 1C), indicating that the observed change in populations and dynamics results in a more polar environment, on average, for the 19F label. While the population change upon addition of GDP-Man (1.7%) as determined from the relaxation dispersion data is significant, given the fitted errors in the analysis of the relaxation data (Table 2), it might not be possible to deconvolute model parameters entirely, given that experiments were acquired at two magnetic field strengths only; however, changes in dynamics and changes in populations toward the less populated state are also manifest in the observed upshift of the inactive state resonance in Fig. 1C, i.e. to the state in which the 19F label is located in a more polar environment. Resonances of 19F nuclei in a more polar environment are located upfield of 19F nuclei buried inside a protein (
). This is in agreement with our observation that the inactive state resonance, for which the 19F label is more exposed to the solvent than for the active state resonance (Fig. 1B), is upfield of the active state resonance. Based on the crystal structures (Fig. 1A) and our MD simulations (Fig. S3), the 19F label is most solvent-exposed in the inactive, extended conformation. Thus, we identify the sparsely populated state with the inactive, extended conformation. This conclusion is further corroborated by structural features of PimA.
Table 2Parameters obtained from CPMG relaxation dispersion experiments of PimAR144C-TFA fitted to Equation 3
The conformational transition from the compact to extended inactive state of PimA, in the absence of GDP-Man, was reproduced by steered-MD simulations, as shown in Fig. S7. This type of calculation forces the system to evolve away from its initial equilibrium condition (compact, inactive) to a final state (extended, inactive), thereby accelerating transitions between different energy minima (
). As revealed by the steered-MD simulations, the inactive, extended state’s large hydrophobic patches are exposed, making this state potentially aggregation-prone, whereas the hydrophobic patches are shielded from solvent in the inactive, compact state (
). The fact that the addition of GDP-Man leads to an increase in population of the inactive, extended state suggests that the slower reshuffling event occurs in that state.
The active state of PimA undergoes millisecond motions that may reflect open-to-closed dynamics
As shown in Fig. 3C, the resonance in the 19F spectrum of PimAR144C-TFA monitoring the active states also displays relaxation dispersion, and the profile is again affected by the addition of saturating amounts of GDP-Man, reflecting changes in dynamics and population equilibria. In contrast to the inactive state resonance, the active state resonance is downshifted upon addition of GDP-Man (Fig. S5). This indicates that a population shift toward a more hydrophobic environment for the 19F label occurs, which might be identified with a population shift from a structurally uncharacterized active, extended state to the active, compact state of PimA. Based on this interpretation, most of PimA is found in the active, extended state (81% extended versus 19% compact). This holds true even after addition of GDP-Man (70% extended versus 30% compact) (Fig. 3D). The active, extended state might be represented by the open version of the already structurally characterized active, closed state, suggesting that these dynamics correspond to the open-to-closed motion of PimA (Fig. 1A). The exchange rate between the extended and compact forms of the active states in this case increases from 4300 s−1 to 8000 s−1 upon addition of saturating concentrations of GDP-Man.
Interestingly, accelerated MD simulations performed on PimA labeled with TFA and starting from the compact, active state indicated that the protein undergoes a conformational change to reach a state in which β7 dissociates from β6, which indicates the initiation of the transition to α-helix (Fig. S8). As steered-MD simulations, this computational approach is useful for improving conformational space sampling. The method does not require any predefined coordinates and reduces the energy barrier between the different low-energy states (
The active and inactive states are structurally distinct
The relaxation dispersion data indicate that the inactive, compact and inactive, extended states are structurally distinct, as their resonances are separated by 1.2 ppm (Table 2). As discussed above, the resonance of the inactive, extended state is expected to be upfield of the resonance of the inactive, compact state, since the 19F label is more solvent-exposed in the latter (Fig. 1C). Similarly, the relaxation dispersion data indicate that the active, compact and active, extended state are structurally distinct, as their resonances are separated by 2.9 ppm (Table 2). The downfield shift of the resonance of the active, compact state compared with the active, extended state (Fig. S5) indicates that in the active, compact state the 19F label is located in a more hydrophobic environment. Importantly, from the relaxation dispersion data (Table 2) and the 1D spectra (Fig. 1C, Fig. S5), it can be concluded that all conformational states are structurally distinct, indicating that fold-switching is accompanied by a dramatic structural rearrangement near the 19F label. This is consistent with the structural data that show that residue Arg144 is reshuffled from an α-helix (potentially via a solvent-exposed random coil structure) to a β-sheet environment.
The ability of an enzyme to adopt many different conformations during the catalytic cycle is due to its great flexibility, playing a critical role in the recruitment of substrates, release of products, stabilization of transition states, and adaptation to the cell environment. Employing 19F NMR, we identified four functionally relevant states of PimA that coexist in dynamic equilibria in solution undergoing conformational exchange on timescales from milliseconds to seconds. Specifically, fold-switching, a hitherto poorly characterized process, occurs in the reshuffling region of the N-terminal domain of PimA on a second timescale. Domain motions, on the other hand, occur on a millisecond timescale.
The observations presented here allow us to propose that the activation of PimA occurs along a sequence of structural rearrangements displayed in Fig. 4. In Fig. S9, a free-energy diagram of the activation process is depicted that is derived from thermodynamic data obtained from the saturation transfer and relaxation dispersion experiments (Table 1, Table 2). Initially, PimA is inactive and in a compact state that shields hydrophobic patches in the reshuffling region. Almost all inactive PimA molecules are in that conformation but are in millisecond exchange with the inactive, extended state. This state exposes hydrophobic patches and, thus, is sparsely populated. However, it offers sufficient degrees of freedom for a rearrangement of secondary structure elements on a second timescale. A drastic change in chemical shift indicates that the inactive, extended state is structurally distinct from the uncharacterized active, extended state, which might correspond to the open state of the structurally characterized closed, GDP-Man-bound conformation of PimA. Following the fold-switch, which we speculate to occur from the inactive, extended state, PimA is in the active, extended state and in millisecond exchange with the active, compact state, which is the conformation in which catalysis occurs. We propose that fold-switching is an activation mechanism of PimA, and since it is a slow process, it may represent the rate-limiting step of the glycosylation reaction catalyzed by PimA. The substrate, GDP-Man, acts as an activator and, thus, functional regulator of PimA, as the binding of GDP-Man enhances the fold-switching dynamics. Intriguingly, a significant population of the inactive state is always present. This may point to a more significant role of the inactive states in the catalytic cycle of PimA than previously anticipated, for instance, in membrane binding of PimA or to facilitate efficient recruitment of phosphatidyl-myo-inositol lipids, which are acceptor substrates of PimA.
This study demonstrates that the 19F-based NMR approach presented here allows for insights into structures and dynamics for proteins that cannot be studied with more conventional NMR methods. The methods employed here, a combination of saturation transfer methods and relaxation dispersion experiments, allow investigations of dynamics occurring simultaneously and on a wide range of timescales in solution. Here, this approach revealed a remarkable situation in which PimA exists in at least four different conformations that are all present in solution. The dynamics between and the relative populations of these conformations are affected by the addition of substrate, which demonstrates that the equilibria are finely tuned to optimize the catalytic cycle.
Materials and Methods
1,1,1-Trifluoroacetone (TFA), 3-bromo-1,1,1-trifluoroacetone (BTFA), guanosine 5′-diphospho-d-mannose (GDP-Man), and tris(2-carboxyethyl)phosphine (TCEP) were acquired from Merck (Darmstadt, Germany).
Cloning and purification of WT PimA and PimA mutants
PimAR144C and PimAT126C-V359C-R144C mutants of PimA from Mycobacterium smegmatis were obtained from GenScript using pET29a-pimA as the DNA template (
). 1 mm TCEP was added to 100–200 μm PimA and incubated for 1 h on ice, except for PimAT126C-V359C-R144C-TFA, to which no TCEP was added. Next, BTFA was added in 10-fold molar excess (1–2 mm), and the solution was incubated overnight at 4 °C under gentle agitation. Finally, the solution was passed through a desalting column equilibrated with 50 mm Tris-HCl, pH 7.5, and 150 mm NaCl. The sample conditions for NMR experiments were 100 μm PimA, 50 mm Tris-HCl, pH 7.5, 150 mm NaCl, and 10% D2O. 500 μm GDP-Man was added where appropriate and if not stated otherwise.
To exchange to 100% D2O solvent, 100 μm PimA was concentrated roughly five times and diluted back to a concentration of 100 μm using 50 mm Tris-HCl, 150 mm NaCl in 100% D2O. This procedure was repeated 5 times.
Experiments were performed at 25 °C on a Bruker Avance 600 MHz III HD spectrometer (19F Larmor frequency of 564 MHz). Relaxation dispersion and titration data for PimAR144C-TFA were additionally acquired on a Bruker Avance 700 MHz spectrometer (19F Larmor frequency of 659 MHz). Both spectrometers were equipped with a 19F cryoprobe. Typical π/2 pulse lengths were 15–19 µs, and the acquisition time was 0.03 s. 10% D2O was added to the samples for frequency locking. For 1D spectra, 400 transients were acquired with a relaxation delay of 1 s. Data were processed in Topspin (Bruker, Billerica, MA) and analyzed using in-house Matlab routines (MathsWorks, Natick, MA) and Igor Pro (WaveMetrics, Lake Oswego, OR), employing an in-house script for nonlinear fitting of relaxation dispersion data (
). Overlapping resonances were deconvoluted, employing iNMR software (MestreLab Research, Spain) by fitting spectra to double-Lorentzian functions.
Far-UV CD analysis
Spectra were acquired in a J-810 CD spectropolarimeter (Jasco Corp., Tokyo, Japan) by using Hellma 105.200-QS quartz cuvettes with a 1 cm optical path. Spectra were recorded in a continuous mode with 1 nm bandwidth, 1 s response, and a scan speed of 100 nm/min−1. Samples were 2.5 μm PimA, PimAR144C, or PimAR144C-TFA in 10 mm Tris-HCl, pH 7.5. GDP was added at a 1:15 ratio, and 25 scans were accumulated to obtain the final spectra, which were further corrected for the baseline signal. Spectra were recorded in the 200–250 nm range at 20 °C.
Spectra were acquired in a J-810 CD spectropolarimeter (Jasco Corp., Tokyo, Japan) by using Hellma 110-QS quartz cuvettes with a 1 mm optical path by using a Peltier thermal device, allowing temperature control during the experiments. Spectra were recorded in a continuous mode with 1 nm bandwidth, 1 s response, and a scan speed of 100 nm/min−1. Samples were 10 μm PimA, PimAR144C, or PimAR144C-TFA in 10 mm Tris-HCl, pH 7.5. Substrates were added at a 1:15 ratio. In this case, thermal dependencies of the ellipticity were monitored in the range from 20 °C to 90 °C at 222 nm. Temperature was increased stepwise by 1 °C/min.
). 2400 transients were acquired during a constant time delay, TCPMG, of 3.84 ms for 14 refocusing frequencies, νCPMG, ranging between ∼130 s−1 and ∼6000 s−1. Effective transverse relaxation rates, R2,eff, were computed from peak intensities I(νCPMG) relative to the peak intensity, I0, in the absence of a constant time delay as follows
For data measured at one magnetic field strength, relaxation dispersion data were fitted to Luz-Meiboom's two-state model to obtain an apparent exchange rate, kapp, of the dynamics (
where , τCPMG is the separation between two 180° pulses and kapp, R2,0, and are parameters of the fit. The populations, pA and pB, of the two states and the chemical shift difference between the resonances, Δω, cannot be deconvoluted with this model.
Relaxation dispersion data measured at two magnetic field strengths were fitted to Carver-Richards model assuming two states to obtain the exchange rate of the process, ki (i indicates intermediate exchange), populations, pA and pB, and the chemical shift difference, Δω, between the states (
To reduce the number of parameters in the fit, it was assumed that the addition of GDP-Man did not alter the conformations of PimA present in solution, i.e. the chemical shift , where f is the spectrometer frequency, was treated as a global constant. Moreover, the magnetic field strength does not affect protein kinetics, i.e. ki, pA, and pB were identical for data obtained at different magnetic field strengths but could vary between samples that contained or did not contain GDP-Man.
Inversion recovery experiments with at least 10 delays, τ, ranging between 25 ms and 800 ms were performed to obtain the longitudinal relaxation time, T1. 1200 transients were acquired, and the relaxation delay was 2 s. The relaxation rate, R1 = T1−1, was obtained from a monoexponential fit of the intensities in dependence on delay τ.
Saturation transfer experiments
Saturation transfer experiments were conducted to quantify slow exchange dynamics of PimA (
). Resonances were saturated with continuous wave irradiation (B1 field of 28 Hz). The resonance associated with the active state was irradiated at −82.3 ppm, and the intensity decay of the inactive state resonance was monitored at −84.1 ppm in the absence of substrate and at −84.3 ppm in the presence of GDP-Man. Conversely, the resonance associated with the inactive state was irradiated at −84.3 ppm, and the intensity decay of the active state resonance was monitored at −82.7 ppm in the absence of substrate and at −82.6 ppm in the presence of GDP-Man. Spectra were acquired for 8 saturation times, varying between 25 ms and 1000 ms. 1200 transients were recorded, and the relaxation delay was 1.5 s. Exchange rates were obtained by fitting residual intensities, , at increasing irradiation times τ to a two-state model (
). Three different simulations, starting from the active-compact, inactive-compact, and inactive-extended conformations, were performed. Parameters for the TFA label were generated with the antechamber module of AMBER implemented with the general Amber force field (GAFF) (
) water molecules. The system was neutralized by adding explicit counter ions (Na+). A two-stage geometry optimization approach was performed. The first stage minimizes only the positions of solvent molecules and ions, and the second stage is an unrestrained minimization of all the atoms in the simulation cell. The systems then were gently heated by incrementally increasing the temperature from 0 to 300 K under a constant pressure of 1 atm and periodic boundary conditions. Harmonic restraints of 30 kcal·mol−1 were applied to the solute, and the Andersen temperature coupling scheme (
) was used to control and equalize the temperature. The time step was kept at 1 fs during the heating stages, allowing potential inhomogeneities to self-adjust. Water molecules are treated with the SHAKE algorithm such that the angle between the hydrogen atoms is kept fixed. Long-range electrostatic effects are modelled using the particle-mesh-Ewald method (
). An 8 Å cutoff was applied to Lennard-Jones and electrostatic interactions. Each system was equilibrated for 2 ns with a 2 fs time step at a constant volume and temperature of 300 K. Production trajectories were then run for an additional 200 ns under the same simulation conditions. SASA for fluorine atoms of the TFA label was calculated using the keyword “SURF” in the cpptraj module (
With the accelerated MD implemented in AMBER 18, 500 ns accelerated MD simulations were performed on PimA labeled with TFA and starting from the active-compact conformation. These simulations were restarted from the final structure of the 200 ns conventional MD simulations with random atomic velocity initializations at 300 K. Boost potential with an extra boost to the torsions was applied (iamd = 3).
The protocol employed above for conventional MD simulations was also used in steered-MD calculations (
). The compact-inactive conformation was used as the starting structure. For the final production trajectory, the distance between C-alpha of Ala148 and Ala366 was varied from 13.5 (close, inactive state, 0 ns) to 51.3 Å (extended, inactive state, 200 ns), using an rk2 of 8 kcal·mol−1·Å−1.
Author contributions—J. L., M. E. G., and L. M. conceptualization; J. L., F. C., and T. S. data curation; J. L. and L. M. software; J. L., F. C., T. S., and L. M. formal analysis; J. L., T. S., M. E. G., and L. M. supervision; J. L., F. C., and T. S. validation; J. L., M. T., B. T., J. P., A. R.-U., F. C., T. S., M. E. G., and L. M. investigation; J. L., M. T., B. T., J. P., A. R.-U., F. C., T. S., and L. M. methodology; J. L., F. C., M. E. G., and L. M. writing-original draft; J. L., F. C., M. E. G., and L. M. writing-review and editing; F. C. and M. E. G. visualization; M. E. G. and L. M. resources; M. E. G. and L. M. funding acquisition; M. E. G. and L. M. project administration.
Funding and additional information—This work was supported by grants from the Swedish Research Council Contract 621-2014-3706 (to L. M.), the MINECO, FEDER-EU contract BFU2016-77427-C2-2-R and contract BFU2017-92223-EXP, the Severo Ochoa Excellence Accreditation SEV-2016-0644 (to M. E. G.) and RTI2018-099592-B-C21 (to F. C.), and the Basque Government contract KK-2019/00076 (to M. E. G.). The Knut and Alice Wallenberg Foundation is acknowledged for financial support for the “NMR for Life” program.
Conflict of interest—The authors declare that they have no conflicts of interest with the contents of this article.
Abbreviations—The abbreviations used are: PIMs
midpoint temperature of unfolding for the unliganded protein