To understand the origin of binding cooperativity in hTS, we first needed to determine if a conformational selection or dynamic allostery mechanism is utilized. In other words, does dUMP binding elicit a change in the structure or the dynamics of hTS, or perhaps both? To begin to answer this question, we needed to establish whether apo hTS, which crystallizes in active and inactive conformations (Chen et al., 2017; Figure 1c), switches between these two conformations in solution. NMR spectroscopy enables the investigation of protein dynamics on a wide range of timescales, including the µs-ms timescale where this type of conformational equilibrium is typically seen. For our characterization of the dynamics of hTS on the µs-ms timescale, we have employed CPMG relaxation dispersion and CEST experiments targeting the methyl groups on isoleucine, leucine, and valine side chains. These experiments probe somewhat overlapping regions of the µs-ms timescale, with CEST characterizing motions on the ms timescale and CPMG characterizing motions with exchange timescales of 100s of µs to 10s of ms. In the CEST experiment, a spin lock is applied at many frequencies scanning the entire spectrum, and the intensity of the observed (major state) signals are monitored to determine frequencies at which saturation is transferred from a minor state to the major state (Vallurupalli et al., 2012). In the CPMG experiment, dephasing of transverse magnetization due to stochastic exchange between states with distinct frequencies (i.e. the contribution of chemical exchange to transverse relaxation, Rex) is refocused using trains of 180° pulses applied at various rates. We have used multiple-quantum (MQ) (Korzhnev et al., 2004) and ¹³C single-quantum (SQ) (Lundström et al., 2007) CPMG experiments as well as a ¹³C CEST experiment using the CHD₂ methyl isotopomer (Rennella et al., 2015). The MQ CPMG and CEST experiments are especially well suited to probing these motions in a large protein like hTS. Our analysis of the µs-ms dynamics of apo hTS include 850 and 600 MHz MQ CPMG datasets, an 850 MHz ¹³C SQ CPMG dataset, as well as 40 and 25 Hz spin lock power CEST datasets. For dUMP bound hTS, the analysis includes 850 and 600 MHz MQ CPMG datasets and an 850 MHz ¹³C SQ CPMG dataset.

The initial analysis of the CPMG and CEST datasets revealed that, while some methyl probes can be well-described as undergoing exchange between two conformations, other probes clearly cannot and appear to be involved in dynamic switching between three states (Appendix 1). Global analysis of the CPMG and CEST datasets (see Materials and methods) on apo hTS show that 10 probes are involved in a concerted process with an exchange rate of 240 ± 7 s^-1 and an excited state population of ~1.3% (Figure 1, Figure 1—figure supplement 1, Supplementary file 2, Figure 1—source data 1 and 2, Figure 2—source data 1–4). Interestingly, 8 of these 10 probes show 3-state exchange (Appendix 1), that in addition to the 2-state concerted process includes a faster process with variable exchange rate and population (Figure 2). Multiple trials of a global fit using a combination of 2- and 3-state models for different methyl sites were performed. For instances of 3-state behavior, a B↔A↔C model, in which each excited state (B and C) exchanges with the ground state (A) but not with each other, is most appropriate in these cases, as a 2-state model does not fit the data well and the A↔B↔C model lacks consistency in the exchange parameters amongst these probes (Appendix 1). These probes involved in the concerted process include L187 and L192 in the active site loop, as well as L198, which lies in the dimer interface on one end of the loop (Figure 1b). In addition, L101, L121, and L131, which lie in the ‘extended loop’, are also involved in this process. Of the remaining four probes, three – L74, V79, and L221 – are around the binding site of the cosubstrate mTHF. The active site and extended loops show significant differences at the backbone level between the active and inactive structures (Figure 1c), with most of the extended loop lacking electron density in the original inactive structure. In the active conformation, the active site loop is oriented towards the substrate binding site, with the catalytically essential C195 making direct contact with dUMP. In the inactive conformation, on the other hand, the active site loop rearranges via a 180° flip (Figure 1c, right) and C195 orients towards the dimer interface. As a result, the interaction between C195 and dUMP that plays a significant role in substrate binding (Gibson et al., 2008; Sapienza and Lee, 2016) is lost. In addition to the changes in the active site and extended loops, there are small displacements of the backbone throughout the molecule (Figure 1c). The fact that these two loops are involved in this concerted process is consistent with this process being exchange between the active and inactive conformations. We have attempted to confirm this by obtaining the chemical shifts of the active and inactive conformations using mutants which stabilize these conformations, but unfortunately these mutants (R163K, M190K, A191K Gibson et al., 2008; Gibson et al., 2008; Luo et al., 2011) either did not express solubly or yielded very poor NMR spectra.

In addition to the probes involved in the concerted process, there are also 11 probes showing 2-state behavior that have variable exchange rates and populations in apo hTS (Figure 2; Supplementary file 2). There appears to be some consistency in exchange rates and populations between nearby probes for these faster motions; for example, L73 and V79 have well-defined exchange rates and minor state populations that are in excellent agreement, as do L41, L252, and L259 (Figure 2c, Supplementary file 2). In cases where the minor state population for the faster motion is well-defined, it is almost always very small (0.11 - 0.24%). The three probes which clearly have a larger minor state population are V58 (1.1 ± 0.2%), I237 (4.3 ± 0.9%), and V285 (5% quantile of Monte Carlo simulations is 2%). Interestingly, these localized faster motions generally have ¹³C Δω’s that are roughly 3 times as large as those of the concerted, slower motion (Supplementary file 2), suggesting considerable change in the local environment of these probes. While the uncertainty in these Δω’s is quite large, even at the lower end of their error bar most of the probes would still have a Δω at least as large as the largest seen in the slower, concerted process (1.4 ppm).

Finally, there are also two probes showing 3-state exchange that do not appear to be involved in the exchange with the inactive conformation (V58 and L259, Supplementary file 2). The parameters for the slower of the two processes for these probes also appear to be distinct from each other.

Global analysis of the CPMG datasets on dUMP-bound hTS (Figure 2—source data 5–8) reveal that the slower, concerted process is lost upon substrate binding, but faster motion is still present. Further, comparison of the ¹³C Δω’s between apo (fast motion) and dUMP-bound states show a significant correlation despite their uncertainty, suggesting that similar configurations are being accessed (Figure 2d). Exchange rates for the faster motion in apo and dUMP-bound states are generally similar as well (Supplementary files 2 and 3). It is difficult to comment on whether these faster motions are suppressed or amplified upon dUMP binding due to the relatively large error in the populations in the dUMP-bound analysis, however the smaller number of analyzable probes in the case of dUMP-bound hTS suggests that these motions are at least somewhat suppressed upon substrate binding (Supplementary file 3). The loss of the slower, concerted process in dUMP-bound hTS is consistent with this process being exchange between the active and inactive conformations, as the inactive conformation binds dUMP with much lower affinity than the active conformation (Chen et al., 2017).

To gain further insights into these faster motions, we performed solvent paramagnetic relaxation enhancement (sPRE) experiments, measuring intensities of the methyl signals in 0 mM and 2 mM gadodiamide (Figure 3—source data 1). Gadodiamide is a paramagnetic cosolute, which increases the relaxation rates (decreases signal intensities) of the methyl signals in a manner dependent on the depth, or the distance to the surface, of the methyl probe in the structure. Interestingly, we find that many of the probes around the active site and in the extended loop showing the faster motion have enhanced sPRE (1 – intensity ratio) and thus are more exposed to the solvent than would be expected given their depth in the ground state structure (Figure 3). Notably, these data were collected on a truncation of hTS lacking the first 25 residues, termed ‘Δ25’, which lacks the slower concerted process but retains the faster localized motions (Appendix 1—figure 2). This enables us to attribute the observed enhancement in solvent exposure to these localized, faster motions as opposed to the active-inactive equilibrium. In addition, we find that probes possessing this unexpectedly large solvent exposure that did not exhibit exchange in our CPMG and CEST global analysis are nearby probes exhibiting the faster motion. We propose that the non-concerted, faster motion involves localized excursions to ‘exposed states’, particularly in regions of the protein surrounding the nucleotide and folate binding sites as well as the extended loop. It should be noted that this enhanced exposure of the active site has been seen in other enzymes (Bernini et al., 2009). In addition to 1D depth, we have also used 3D atom depth index (Varrazzo et al., 2005) for the analysis, yielding very similar results (Figure 3—figure supplement 1). Exceptions to this are L67, L198, L279, and V285, which possess the faster motion but do not show enhanced solvent exposure. These probes are marked with asterisks in Figure 3. L279 and V285 lie on the opposite end of each protomer relative to the substrate binding site, while L67 and L198 resides at the dimer interface. We note that the lack of enhanced solvent exposure at probes on the dimer interface (Figure 3b), the lack of concertedness of the faster motion (Figure 2c), and the presence of the faster motion in the dUMP bound enzyme all indicate that this faster motion is not exchange between the dimer and monomeric forms of hTS. The observed enhancement in solvent exposure is consistent with the characteristically large ¹³C Δω’s of the faster motion, as this would require relatively large structural changes.

The CPMG and CEST data show that apo hTS undergoes exchange between the active and inactive conformations, and that this exchange process is coupled to dUMP binding. Yet the question remains as to which of these conformations is the ground state (~98.5%) and which is the excited state (~1.3%). Only if the inactive conformation is the ground state can a conformational selection model contribute significantly to the binding cooperativity, assuming that the inactive conformation binds dUMP much more weakly than the active conformation.

In order to determine which conformation is dominant in solution in apo hTS, we measured residual dipolar couplings (RDCs) on the backbone amides (Figure 4, Figure 4—source data 1 and 2). These RDCs contain information regarding the orientation of the amide bond vectors; thus, by comparing the experimentally observed RDCs with those calculated from the active and inactive structures using a fit alignment tensor (see Materials and Methods), it can be determined which of the two conformations, if either, is consistent with the data. As a comparison, we also measured RDCs on the backbone amides of dUMP-bound hTS (Figure 4—source data 3), where the ground state structure is known. The comparisons show that the active conformation is in significantly superior agreement with the apo hTS data than the inactive conformation (Figure 4). Further, we find that the agreement between the apo RDCs and the active structure is just as good as the agreement between dUMP- bound RDCs and the hTS-dUMP structure (also active), having essentially identical correlation coefficients (0.91 and 0.92, respectively). Together, these findings demonstrate that the active conformation is the ground state of apo hTS, indicating that the minor state of the concerted process observed in the CPMG and CEST data is the inactive conformation (1.3%). Consistent with this notion, we find that changes in ¹³C chemical shift upon dUMP binding correlate very poorly with, and are generally much smaller in magnitude than, the ¹³C ∆ω’s obtained for the concerted process (Figure 4—figure supplement 1). The small population of the inactive conformation in the apo enzyme means that conformational selection contributes negligibly to the substrate binding cooperativity. Since the apo active and dUMP-bound structures are so similar (all atom RMSD = 0.84 Å between 5X5 A and 5X5 D structures), this indicates that there is no significant structural change to the enzyme upon dUMP binding. Consequently, the origin of the dUMP binding cooperativity is likely related to the dynamics of the enzyme.

Given that we have excluded conformational selection as the source of dUMP binding cooperativity, ps-ns timescale dynamics were considered since these motions are known to impact binding energetics (Caro et al., 2017; Kasinath et al., 2013). The dynamics of hTS on this much faster timescale were investigated by measuring ²H transverse relaxation rates of CHD₂ methyl isotopomers in isoleucine, leucine, and valine side chains (Tugarinov et al., 2005). From these relaxation rates, we calculate the methyl symmetry axis order parameter, Saxis2 , which takes on a value between 0 and 1. A value of 0 indicates maximal flexibility of the methyl group, while a value of 1 indicates maximal rigidity. These order parameters are related to the conformational entropy of the molecule; thus, changes in the order parameter upon substrate binding are related to the conformational entropy component of the binding entropy. Reasonable estimates of the change in conformational entropy can be made from these order parameter changes using the ‘entropy meter’ (Caro et al., 2017). This approach assumes that the changes in ps-ns timescale motion seen in the isoleucine, leucine, and valine side chains are representative of all the residues in the protein, and that the overall change in conformational entropy is dominated by the side chains.

We first measured the ²H transverse relaxation rates for apo and dUMP bound hTS (Figure 5—source data 1–2). The average change in order parameter (dUMP-apo) is 0.027 ± 0.006 (n = 47) (Figure 5, Supplementary files 7 and 8). Using the entropy meter (see Materials and methods), this corresponds to a -T∆Sconf of 10 ± 3 kcal/mol. The magnitude of this entropic change is about twice as large as the 4.9 ± 0.3 kcal/mol difference between the entropies of the two dUMP binding events (Bonin et al., 2019). This means that if most of the rigidification occurs upon the first binding event, this change in the ps-ns timescale dynamics could be the origin of the positive binding cooperativity.

Probes showing the most rigidification (largest increase in order parameter) are L85, L88, L89, L121, V158, L192, L198, and L221 (Figure 5c). Interestingly, much of this rigidification occurs around the binding site of the cosubstrate mTHF (L85, L88, L89, L221). Notably, our ps-ns dynamics analysis lacks probes in the dUMP binding site (Figure 5b). In addition, there appears to be considerable colocalization of probes which are rigidified on the µs-ms timescale (i.e. probes which lose the exchange with the inactive conformation) and probes which are rigidified on the ps-ns timescale (Figure 6). Of the 6 probes involved in the active-inactive switching (10 total) that are present in the ps-ns dynamics analysis, 4 (L121, L192, L198, L221) show large changes in order parameter (|ΔSaxis2|> 0.06). The remaining 2 probes (L187, I237), while not displaying a large change themselves, are each in close proximity to a residue possessing a large change in order parameter (V158 and L233, respectively, magenta ovals in Figure 6). In addition, there are three probes which have a large change in order parameter that are not in the immediate vicinity of residues involved in the slow motion (L85, L88, and L89, magenta asterisks in Figure 6). However, ¹H 2-plane CPMG data clearly show that V84, and to a lesser extent L85, is dynamic on the µs-ms timescale in apo hTS (Figure 6—figure supplement 1). Further, the helix containing these residues (residues 80–93, referred to hereafter as the folate binding helix) connects probes involved in the concerted process, specifically L101 and V79. These observations suggest that L85, L88, and L89 may be involved in the slow motion as well. Finally, 8 of the 9 probes with a large change in order parameter (red/purple in Figure 6) showed an increase in order parameter upon dUMP binding (more rigid), the exception being L233.

To further establish the connection between the ps-ns timescale dynamics and the binding cooperativity, we have also investigated these dynamics in two very similar scenarios which lack binding cooperativity. The first is binding of the product TMP to hTS. TMP is very similar to the substrate dUMP, differing only by the addition of a methyl group on the nitrogenous base, yet we have found that hTS binds TMP non-cooperatively (Appendix 1). Further, chemical shift perturbations (CSPs) between the TMP and dUMP-bound forms are generally of a similar (small) magnitude as the CSPs between the dUMP-bound and apo forms (Figure 7a and b), suggesting that there is also little structural change between hTS-TMP and hTS-dUMP (there is currently no crystal structure of hTS-TMP). Interestingly, many of the probes having the largest CSPs are clustered at the dimer interface, which is consistent with there being a difference in the intersubunit communication between the TMP and dUMP-bound forms. These probes include L198 and L212 on the beta sheet, as well as L187 from the active site loop and V158 from another loop.

The small CSPs between hTS-TMP and hTS-dUMP again point to dynamics as being the driving force behind the dUMP binding cooperativity. We have measured ²H relaxation rates for hTS-TMP as well (Figure 5—source data 3), and we find that the average change in order parameter between hTS-TMP and apo hTS is -0.006 ± 0.006 (n = 46) (Figure 5, Supplementary files 7 and 9). This corresponds to a -T∆Sconf of -2 ± 2 kcal/mol, meaning that in this case there is actually a slight increase in flexibility of the molecule, as opposed to the rigidification observed with dUMP binding.

The second scenario we consider is dUMP binding to Δ25 hTS. As in the case of TMP binding, this truncated form of hTS binds dUMP non-cooperatively (Appendix 1—figure 3), a fascinating result given that the N-terminus is absent in crystal structures of both apo and dUMP bound hTS. It is notable that the non-cooperative E. coli enzyme lacks this N-terminal extension, identifying it as a region of interest as a possible determinant of the cooperativity. Δ25 hTS is perhaps an even more striking example than hTS-TMP, as there are essentially no perturbations of the chemical shifts at all to the apo enzyme upon removal of the N-terminus (Figure 7c), suggesting that the structures are nearly identical. This suggests once again that differences in the dynamics of these molecules are behind the differences in binding cooperativity.

Consequently, we have measured ²H transverse relaxation rates for apo and dUMP-bound Δ25 hTS as well (Figure 5—source data 4–5). We find that for Δ25 hTS binding dUMP, the average change in order parameter is 0.009 ± 0.004 (n = 76) (Figure 5, Supplementary files 10 and 11), corresponding to a -T∆Sconf of 3 ± 2 kcal/mol. While there is still a slight rigidification in this case, the magnitude of the entropic change is less than 1/3 that of dUMP binding to full length hTS. It is worth noting that our analysis of dUMP binding to full length hTS includes only a single residue from the N-terminus itself, V3, as the two N-terminal leucines (L8 and L13) are unassigned. This is to say that the observed differences between full length and Δ25 hTS stem from the N-terminus’ impact on the rest of the protein, rather than from the N-terminus itself. These results establish a correlation between the rigidification upon dUMP binding and cooperativity.

Since the rigidification on the ps-ns timescale upon dUMP binding is likely the source of dUMP binding cooperativity, we sought to determine the origins of this rigidification. An answer is found in considering a thermodynamic cycle consisting of dUMP binding to the full length and Δ25 enzymes (Figure 8). We define two paths, A and B, which both begin and end with full length apo and full length dUMP-bound, respectively. Path A contains a single step, which is dUMP binding to the full length enzyme. Path B contains three steps: (1) removal of the N-terminus from the apo enzyme, (2) dUMP binding to the truncated enzyme, and (3) addition of the N-terminus to the truncated dUMP-bound enzyme. The changes in conformational entropy for each of these steps is calculated based on the order parameter values discussed previously. We find that the summed change in conformational entropy across each of these paths agree well with each other, as expected. We also find that the change in conformational entropy upon removal of the N-terminus from the apo enzyme (step 1 of path B) is larger in magnitude than the change in conformational entropy upon addition of the N-terminus to the truncated dUMP bound enzyme (step 3 of path B). This means that the N-terminus has a larger impact on the ps-ns timescale dynamics of the apo enzyme than the dUMP-bound enzyme, suggesting that there are interactions between the N-terminus and the protein surface in apo hTS that are coupled to dUMP binding. These interactions make the apo enzyme more flexible (step 1 of path B has a positive sign). Further, we find that the conformational entropy change associated with this coupling (step 1+step 3 of path B or equivalently path A – step 2 of path B, 6 ± 3 kcal/mol when calculated using probes common to all states) is larger than the conformational entropy change of dUMP binding to the truncated enzyme (step 2 of path B, 3 ± 2 kcal/mol), making this coupling of the N-terminal interactions responsible for the majority of the conformational entropy change on path B. While the uncertainties in the conformational entropy change at each step are fairly large, the change in conformational entropy along path A is clearly larger than step 2 of path B, demonstrating that the N-terminus plays a noticeable role in this change. Together, these results identify the N-terminus of hTS as a key player in the rigidification of the enzyme on the ps-ns timescale upon dUMP binding, suggesting that interactions between the N-terminus and the protein surface, which are diminished/altered upon dUMP binding, contribute significantly to the overall change in conformational entropy. Ironically, it is this flexible N-terminus (which lacks electron density in all crystal structures of hTS) which enables greater rigidification upon dUMP binding.

To gain a deeper understanding of how the N-terminus affects the fast timescale dynamics of hTS, we turn our attention to the differences in order parameter changes between [apo Δ25 and apo full length hTS] and [dUMP-bound Δ25 and dUMP-bound full length hTS] (step 1+step 3 of path B). The location of these differences in ∆Saxis2 , or ∆∆Saxis2 , are shown on the hTS structure in (Figure 8b). We find that the probes with the largest ∆∆Saxis2 are L121, V158, and L233. ∆∆Saxis2 values for these probes are nearly identical to their ∆Saxis2 values upon dUMP binding to full length hTS, indicating that the entirety of their ps-ns dynamic response to substrate binding is due to the N-terminus (i.e. the change in order parameter of these probes in step 2 of path B is close to 0). Interestingly, the ∆∆Saxis2 values for these probes contain sizable contributions from both the apo and dUMP bound terms, highlighting that despite the larger magnitude of entropy change for step 1 compared to step 3 of path B, the N-terminus still has some effect on the dUMP bound state (i.e. the entropy change of step 3 of path B is nonzero). Additionally, we find that L192 and L221, which are significantly rigidified upon dUMP binding to full length hTS, have ΔΔSaxis2≅ 0. This suggests that the entirety of the rigidification of these probes, which lie near the substrate binding site, is due to the direct interactions between dUMP and hTS.

Samples of hTS were expressed and purified as described previously (Bonin and Lee, 2021), with modifications made for the specific methyl labeling utilized here. Briefly, a pET21a vector including hTS with an N-terminal 6 x histidine tag was transformed into BL21 (DE3) cells lacking the rne131 gene (Invitrogen, Carlsbad, CA). After an adaptation period (Bonin and Lee, 2021), the cells were grown in M9 media made with D₂O (99.8%, Sigma-Aldrich, St. Louis, MO/Cambridge Isotope Laboratories, Inc, Tewksbury, MA), 1 g/L ¹⁵NH₄Cl (Sigma-Aldrich, St. Louis, MO/Cambridge Isotope Laboratories, Inc, Tewksbury, MA) and 2 g/L U-[²H] glucose (Sigma-Aldrich, St. Louis, MO/Cambridge Isotope Laboratories, Inc, Tewksbury, MA). Cells were grown up to an OD₆₀₀ of 0.65 at 37 °C, at which time one of two Ile and Leu/Val precursors was added: 60 mg/L 2-ketobutyric acid-4-¹³C,3,3-d₂ sodium salt hydrate (Ile) and 100 mg/L 2-keto-3-(methyl-d₃)-butyric acid-4-¹³C, 3-d sodium salt (Leu/Val) for {I (¹³C_δH₃), L(¹³C_δH₃,¹²C_δD₃), V(¹³C_γH₃,¹²C_γD₃)} U-[²H,¹⁵N] hTS (referred to hereafter as ILV CH₃ hTS) or 60 mg/L alpha-ketobutyric acid-4-¹³C,3,3,4,4-d₄ sodium salt (Ile) and 100 mg/L alpha-ketoisovaleric acid-3-methyl-¹³C,3-methyl-d₂,3,4,4,4-d₄ sodium salt (Leu/Val) for {I (¹³C_δHD₂), L(¹³C_δHD₂,¹²C_δD₃), V(¹³C_γHD₂,¹²C_γD₃)} U-[²H,¹⁵N] hTS (referred to hereafter as ILV CHD₂ hTS). One hour after the addition of the precursors, IPTG was added to 1 mM, and protein was expressed at 20 °C for 12 hr. For samples without the ILV labeling (U-[²H,¹⁵N] hTS, referred to hereafter as ¹⁵N hTS), IPTG is simply added at an OD₆₀₀ of 1.0, followed by expression at 20 °C.

The purification workflow was as described previously (Bonin and Lee, 2021). Notably, cleavage of the 6 x histidine tag leaves an extra glycine and alanine at the N-terminus. Protein was concentrated to ~0.2 mM (dimer) for NMR study, as determined by 280 nm absorbance using ε280=117,040Lmol∗cm (Bonin et al., 2019). The final conditions for NMR study were 27.5 mM Na₂HPO₄, 20 mM NaCl, 0.02% NaN₃, 5 mM DTT, pH 7.5 (NMR buffer) with D₂O added to 5%, and a temperature of 25 °C. For samples in D₂O, the NMR buffer was prepared in 99.8% D₂O and pD (uncorrected) was 7.1 (Covington et al., 1968). The sample was exchanged into the D₂O buffer using a NAP-5 Sephadex G-25 column (GE Healthcare, Chicago, IL). For dUMP and TMP (Sigma-Aldrich, St. Louis, MO) bound samples, nucleotide was added to a ratio of ~10:1 nucleotide:dimer as determined by 260 nm absorbance using ε260=9660Lmol*cm (dUMP) or ε260=7936Lmol*cm (TMP).

A 0.2 mM sample of apo ¹⁵N hTS in NMR buffer (5% D₂O) was prepared as described. Residual dipolar couplings (RDCs) of backbone amides were determined using the ARTSY method (Fitzkee and Bax, 2010). First, a measurement of the isotropic coupling (J coupling) was made, after which Pf1 phage (ASLA Biotech, Riga, Latvia) was added to 7 mg/mL, resulting in a 4.7 Hz splitting of the D₂O line. A second coupling measurement was made in this aligned condition (J+RDC), and the RDC was calculated by taking the difference between aligned and isotropic couplings. dUMP was then added to the aligned sample to 2.5 mM, and the aligned coupling measurement was repeated. RDCs for dUMP bound hTS were computed using the J couplings from apo hTS. The error in signal intensities was taken to be 1.5 x base plane noise based on the duplicate point RMSD observed in pseudo-3D datasets. These measurements were made on a Bruker Avance III HD 850 MHz spectrometer with a TCI H-C/N-D 5 mm cryoprobe. Additional details regarding acquisition parameters for these and all other NMR experiments can be found in Supplementary file 5. Unless otherwise stated, all NMR data were processed using NMRPipe (Delaglio et al., 1995) and signal intensities were obtained using NMRViewJ (Johnson and Blevins, 1994).

RDC data were analyzed using the DC web server. The alignment tensor was fit (order matrix method) using the observed RDCs and a structure of hTS (5X5A: active, 1YPV: inactive, 5X5D: dUMP bound), with the calculated RDCs being computed from the alignment tensor and the structure. Residues from the flexible N and C termini (1-29, 301-313) were excluded from this analysis, as were residues 108–129, which lack electron density in the inactive structure.

For MQ CPMG (Korzhnev et al., 2004) experiments, a 0.2 mM sample of LV CH₃ apo hTS in NMR buffer (99.8% D₂O) was prepared as described. Datasets were collected at 850 and 600 MHz. dUMP was added to 2.2 mM and datasets were recollected. For ¹³C SQ CPMG (Lundström et al., 2007) experiments, a 0.3 mM sample of ILV CH₃ apo hTS in NMR buffer (99.8% D₂O) was prepared as described. Datasets were collected at 850 MHz with ¹³C carrier frequencies at 24 ppm (Leu/Val) and 14 ppm (Ile). dUMP was added to 2.8 mM and datasets were recollected. For ¹³C CEST (Rennella et al., 2015) experiments, a 0.3 mM sample of ILV CHD₂ apo hTS in NMR buffer (99.8% D₂O) was prepared as described. Datasets were collected at 600 MHz using spin lock powers of 41 and 25 Hz as determined by calibration via nutation experiment. 500 Hz ²H decoupling was applied during the relaxation period.

For CPMG datasets, R2,eff values were calculated from peak intensities (Source code 1) using the following equation:R2,eff(planei)=−1Trelaxln⁡(Int(planei)Int(refplane))

where Trelax is the relaxation time and plane i is the ith plane with 180° pulses applied at frequency υcpmg(i). The error in signal intensity was taken to be the duplicate point RMSD for the CPMG datasets and 1.5 x base plane noise for the CEST datasets. The three sets of R2,eff values (MQ 850 MHz, MQ 600 MHz, and ¹³C SQ 850 MHz), along with two sets of normalized intensities from the CEST experiments (41 and 25 Hz spin lock) were globally analyzed for each probe (for dUMP bound hTS, the analysis includes only the CPMG data). Initially, data were fit to a 2-state model (Source code 8 and 10). For some probes, however, the data were not well described by a 2-state model, and a bifurcated 3-state model (B↔A↔C, Source code 2–9) was employed in which the A↔B process was concerted. The R2,eff and normalized intensity values are calculated from the exchange parameters using the Bloch-McConnell equations (see Appendix 1 for relevant matrices). For a detailed consideration of alternative 3-state models, see Appendix 1. For the 3-state model fits, individual rates were fit as opposed to exchange rates and populations (kab , kba , kac , kca). The reason for this is that when there is 3-state exchange where only one of the two processes is concerted, only the rates (and not the populations) are identical amongst the probes involved in the concerted process. For probes involved in the concerted process, kab , kba , and Δω_C,ab were fixed as values obtained from a global 2-state fit of the CEST data alone (Source code 11–13 ), as 3 (2)-state fits of the CPMG and CEST data without any fixed parameters were found to give the same answer. The one exception to this is I237, whose Δω_C,ab was found to change noticeably in a 3-state fit of the CPMG and CEST data where no parameters were fixed. ¹H Δω’s were fixed at 0 ppm unless a nonzero value was required to obtain a reasonable fit to the data (Korzhnev et al., 2004). Geminal methyls of the same residue were fit globally. The errors in fit parameter values are taken as the standard deviation of the values obtained in fits of 200 Monte Carlo simulations of the data.

A 0.1 mM sample of LV CH₃ apo Δ25 hTS in NMR buffer (99.8% D₂O) was prepared as described. HMQC spectra were recorded with 0 and 2 mM gadodiamide (APExBIO, Houston, TX). These spectra were collected on an 850 MHz spectrometer. The intensity ratio is defined as:IntRatio= Int(2mMGd)Int(0mMGd)

The error in signal intensity was taken to be 1.5 x base plane noise. Residue depths were calculated from the 5X5A structure using EDTSurf (Xu et al., 2013).

A 0.2 mM sample of ILV CHD₂ apo hTS in NMR buffer (5% D₂O) was prepared as described. All ²H R2 relaxation (Tugarinov et al., 2005) data were collected on a 600 MHz spectrometer. In the absence of duplicate points (Supplementary file 5), the error in peak intensity was estimated as 1.5 x the base plane noise. dUMP was added to 2 mM to the apo sample, and ²H relaxation data were collected as for apo. For the product-bound state, a 0.2 mM sample of ILV CHD₂ hTS-TMP in NMR buffer (5% D₂O, 2 mM TMP) was prepared as described. ²H relaxation data were collected as for apo and dUMP bound forms. For the truncated enzyme, a 0.2 mM sample of ILV CHD₂ apo Δ25 hTS in NMR buffer (99.8% D₂O) was prepared as described. After collection of the data on the apo enzyme, dUMP was added to 1.7 mM and the data were recollected.

Peak intensities were normalized relative to the first point (Trelax = 0 ms) and fit to a monoexponential decay to obtain the relaxation rate, R2,2H. The errors in transverse relaxation rates were taken as the standard deviation of the values obtained in fits of 200 Monte Carlo simulations of the data (derived from errors estimated from duplicate points or baseplane noise). The ²H relaxation rate, along with the rotational correlation time, τc, of the molecule (see below), was then used to calculate the methyl rotation axis order parameter, Saxis2 , using the following equation:R2,2H=180e2qQℏ2Saxis2τc

where e2qQh is the quadrupolar coupling constant with a value of 167 kHz (Tugarinov et al., 2005).

Changes in conformational entropy were calculated from changes in the methyl symmetry axis order parameter using the following equation:-T∆Sconf=-298.15*sd∆Saxis2Nχ

where ΔSaxis2 is the average change in order parameter, Nχ is the number of side-chain dihedral angles in the protein (1136 used here, calculated using residues 26–307), and sd=-0.0011kcalmol*K is the slope of the entropy meter (Caro et al., 2017). Only probes with errors in ΔSaxis2< 0.07 were included in the analysis.

Samples of 0.1 mM ILV CH₃ apo hTS and 0.2 mM LV CH₃ Δ25 apo hTS in NMR buffer (99.8% D₂O) were prepared as described. HMQC (Tugarinov et al., 2003) spectra were collected on an 850 MHz spectrometer.

Rotational correlation times for the various states of hTS were determined by ¹⁵N relaxation (Lakomek et al., 2012) collected on samples of closely matched concentration. For the apo form, data were collected on separate samples at two fields, 600 MHz [500 MHz]. A 0.2 [0.2] mM sample of ¹⁵N apo hTS in NMR buffer (5% D₂O) was prepared as described. T1 , T1ρ , and heteronuclear NOE datasets were collected at both fields. All T1ρ datasets, unless otherwise stated, were collected with 1.5 kHz spin lock power as determined by nutation experiment. For the dUMP bound form, T1 and T1ρ datasets were collected at two fields, 600 MHz [500 MHz], and a single heteronuclear NOE dataset was collected at 600 MHz. A 0.25 [0.25] mM sample of ¹⁵N hTS-dUMP (2.5 mM dUMP) in NMR buffer (5% D₂O) was prepared as described. For the TMP bound form, T1 , T1ρ , and heteronuclear NOE datasets were collected at 600 MHz. A 0.2 mM sample of ¹⁵N hTS-TMP (2.8 mM TMP) in NMR buffer (5% D₂O) was prepared as described. The T1ρ dataset was collected with 2 kHz spin lock power as determined by nutation experiment. For the apo form of Δ25 hTS, T1 , T1ρ , and heteronuclear NOE datasets were collected at 600 MHz. A 0.2 mM sample of ¹⁵N apo Δ25 hTS in NMR buffer (5% D₂O) was prepared as described. For the dUMP bound form of Δ25 hTS, T1 and T1ρ datasets were collected at 600 MHz. A 0.2 mM sample of ¹⁵N Δ25 hTS-dUMP (2 mM dUMP) in NMR buffer (5% D₂O) was prepared as described.

Intensities from T1 datasets were normalized relative to the Trelax= 0 s point and fit to a monoexponential decay to obtain the longitudinal relaxation rate, R1. Intensities from T1ρ datasets were normalized relative to the Trelax= 0 s point and fit to a monoexponential decay to obtain R1ρ. Transverse relaxation rates, R2, were then calculated from R1ρ, R1, and tilt angle θ using the equation:R2=R1ρsin2⁡θ-R1tan2⁡θ

The heteronuclear NOE was calculated as Int(saturated)Int(ref). Error in signal intensity was taken to be the mean residual of the best fit for T1 and T1ρ datasets, and 1 x base plane noise for heteronuclear NOE datasets. The ROTDIF web server (Berlin et al., 2013) was used to fit τc values for these different states of hTS given the relaxation rates and the 5X5A (apo) or 5X5D (dUMP, TMP) structure. The criteria for inclusion of residues in this fit were as follows: R1 and R2 values were subject to log₂ transformation to normalize their distributions, after which the mean and standard deviation of the normalized distributions were calculated (residues from flexible N- and C-termini were not included in these calculations). Residues with log2⁡R1 and/or log2⁡R2 values outside one standard deviation of the mean were excluded from the analysis. In addition, in cases where NOE data were present, any residue with an NOE <0.65 were excluded from the analysis. In the case of apo Δ25 hTS, residues with NOE <0.6, as well as residues identified by ROTDIF as possessing Rex, were excluded from the analysis. Notably, specific criteria for which residues to include, as well as the type of diffusion tensor used (isotropic, axially symmetric, fully anisotropic) had very little impact on the fit τc,eff . Fits to axially symmetric and fully anisotropic models showed only slight anisotropy to the diffusion tensor, justifying use of the isotropic model for analysis of the ²H relaxation data. Note that this rotational correlation time is determined in 5% D₂O; for samples in 99.8% D₂O, the correlation time is corrected based on the relative viscosities of D₂O and H₂O:τc,D2O=1.235*τc,5% D2O0.95+0.05*1.235

where τc,D2O and τc,5% D2O are the rotational correlation times in 99.8% and 5% D₂O, respectively, and 1.235 is the ratio of the viscosities of D₂O and H₂O at 25 °C (Baker, 1936). Rotational correlation times for all states can be found in Supplementary file 6.