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Protein photosensors provide versatile tools to study ligand-regulated allostery and signaling. Fundamental to these processes is the amount of energy that can be provided by a photosensor to control downstream signaling events. Such regulation is exemplified by the phototropins, plant serine/threonine kinases that are activated by blue light via conserved LOV (Light, Oxygen and Voltage) domains. The core photosensor of oat phototropin 1 is a LOV domain that interacts in light-dependent fashion with an adjacent α-helix (Jα) to control kinase activity. We used solution NMR measurements to quantify the free energy of the LOV domain:Jα helix binding equilibrium in the dark and lit states. These data indicate that light shifts this equilibrium by ~3.8 kcal mol−1, quantifying the energy available through LOV-Jα for light-driven allosteric regulation. This study provides insight into the energetics of light sensing by phototropins and benchmark values for engineering photoswitchable systems based on the LOV-Jα interaction.
Information transfer in biological signaling pathways often takes the form of stimulus-induced changes in protein structure that consequently alter a functional output. Many signaling proteins are composed of linked arrays of modular domains, which cooperatively function to control activity1,2. In such proteins, sensor domains receive inputs related to changes in the local environment, e.g. the binding of a metabolite or other protein, or the alteration of a bound cofactor by a change in redox state. These events change the interactions of the sensors with other domains, which subsequently transmit the signal downstream through changes in their activities. A critical parameter in the construction of such sensors is the magnitude of input-induced changes in the energetics of the sensor domain and its interactions with downstream effectors. It is this change in energy, and the corresponding change in interaction equilibria, that determines the dynamic range of the sensor along with several other fundamental signaling properties.
Several examples of this principle are provided by photosensory proteins, which have evolved to sense and respond to light across the UV/visible spectrum at a wide range of intensities3. These sensory processes are typically achieved through protein domains that bind light-absorbing chromophores and convert photon energy into structural, dynamic and functional changes. One class of these proteins are phototropins, a group of blue light-activated serine/threonine kinases that control a range of biological responses in algae and plants, including phototropism, chloroplast migration and stomatal opening4. Phototropins sense light through two LOV (Light-Oxygen-Voltage) domains, LOV1 and LOV2, which are located on the N-terminal side of the conserved kinase domain (Fig. 1a)5. LOV domains are a subgroup of the larger PAS (Per-ARNT-Sim) domain family, whose members function as sensors of a wide variety of environmental processes throughout biology6,7. To sensitize phototropin to blue light, both LOV1 and LOV2 contain non-covalently bound FMN chromophores (Fig. 1a)8. Blue light absorption induces formation of a covalent adduct between a conserved cysteine residue in the LOV domain and the C4a carbon of the isoalloxazine ring of FMN9. While the functional significance of light absorption by LOV1 is still unclear, adduct formation in LOV2 is crucial for light-dependent enhancement of phototropin kinase activity as assessed in vitro by autophosphorylation and in planta by phototropism and other responses10. Crystal structures and high-resolution solution NMR data of several LOV domains11–16 show they all adopt typical PAS domain folds, consisting of a mixed α/β fold of approximately 110 amino acid residues surrounding the flavin chromophore. Solution NMR studies13 and a subsequent crystal structure15 of a fragment of the Avena sativa phototropin 1 (AsPhot1) containing the LOV2 domain plus a 40 residue C terminal extension have revealed an additional 20 residue α-helix, Jα, that packs onto the PAS core in the dark state. As demonstrated by NMR and other biophysical data, light induces conformational changes in the LOV2 domain that lead to Jα unfolding and dissociation from the LOV domain after the Cys-FMN adduct is formed13,17. Complementary biochemical studies have shown that destabilization of the LOV2-Jα interaction by mutagenesis activates full-length Arabidopsis thaliana phototropin 1 independent of light18. Together, these data argue that dissociation of Jα – or more precisely, a shift in the LOV2-Jα binding equilibrium to favor the helix-dissociated state – upon photon absorption is the principal mechanism by which phototropin is controlled by light (Fig. 1b).
Despite substantial progress in understanding the structural mechanisms of phototropin signaling, the energetics of this core LOV2-Jα switch have not yet been explored. Here we use NMR spectroscopy to quantify the equilibria between the helix-bound and helix-dissociated conformations of LOV2-Jα in the dark and lit states, yielding an estimate of the energy that is potentially available in this photoswitch to achieve this change in functional states. Using NMR 15N and 13C relaxation dispersion analyses, we found that the Jα helix and its binding surface on LOV2 undergo substantial fluctuations on micro- to millisecond timescales in the dark. Fitting these dispersion data to a model for two-site exchange, we determined that the populations of the low- and high-energy conformations are 98.4% and 1.6%, respectively, with a corresponding free energy difference of 2.4 kcal mol−1 across this equilibrium. By comparing the fitted chemical shifts of the high energy state to those directly measured in an isolated Jα peptide, we suggest that Jα is unfolded and dissociated in the high energy conformation despite the lack of a Cys-FMN adduct. Systematic differences between the chemical shifts of the Jα peptide and those of a photoactivated sample suggest that the LOV2-Jα system samples an analogous bound-dissociated equilibrium for Jα after blue-light absorption. A quantitative analysis of these differences indicates that the population of the helix-dissociated conformation in the lit state is ~91%, corresponding to approximately –1.4 kcal mol−1 of free energy across the equilibrium. Thus, we determined that the primary effect of light absorption and the corresponding Cys-FMN adduct formation is to shift the LOV2-Jα binding equilibrium by ~3.8 kcal mol−1. This provides the first estimate of the energy potentially available through the LOV2-Jα photoswitch to activate the kinase domain in AsPhot1, and also yields a critical value for the engineering of artificial photoswitches where the LOV2-Jα interaction is coupled to regulation of the activity of other proteins.
Conformational exchange processes occurring on micro- to millisecond timescales contribute to the transverse relaxation rate (R2) of nuclear spins in NMR spectroscopy. This additional contribution to R2 is termed Rex and can be suppressed by a Carr-Purcell-Meiboom-Gill (CPMG) spin echo pulse train sequence, provided that the exchange process has a similar frequency as the applied 180-degree pulses in the CPMG train (νCPMG; effectively 100–1000 s−1). By examining the effective relaxation rate (R2eff) as a function of νCPMG, one obtains a relaxation dispersion curve that contains thermodynamic, kinetic and structural information on the underlying exchange process19,20. For two-site exchange between a low energy conformation A and high energy conformation B, we define kex as the sum of the forward and reverse reaction rates, pA and pB as population of the two states and Δω as the chemical shift difference between them. Under favorable conditions, these parameters can be extracted from the relaxation dispersion curve.
We used CPMG-based relaxation dispersion experiments on 15N-labeled backbone amides as well as 13C-labeled sidechain methyl groups to probe µs-ms conformational dynamics in the LOV2-Jα protein. Representative dispersion curves of K534 15N backbone amide (Fig. 2a) and L514 13Cδ1 methyl (Fig. 2b) show significant variation in R2eff as νCPMG was increased from approximately 100 to 900 Hz, providing clear evidence for motions on the µs-ms timescale at these two sites. Complete analysis of these relaxation dispersion data revealed that 25 sites exhibited substantial amide 15N and methyl 13C Rex (> 4.0 s−1 and > 2.0 s−1, respectively). Mapping these residues onto the LOV2-Jα structure (Fig. 2c for 15N; Fig. 2d for 13C), we observed that residues with high Rex contributions were clustered on the Jα helix, the IJ loop that connects it back to LOV2 and the β-sheet of LOV2 that faces Jα. Global fits of all 15N and 13C dispersion curves to a model for two-site exchange converged to yield kex = 1320 ± 36 s−1 and pB = 1.6 ± 0.03%. The convergence of the fitting indicated that more complex exchange models were not needed to describe the data, and that the system is reasonably represented as a rapid two-state equilibrium between low and high energy conformations.
Since the majority of resonances with large Rex values were located at the LOV2- Jα interface, we speculated that the dynamics we observed by NMR might involve fluctuations between helix-bound and helix-dissociated conformations. To test this hypothesis, we examined the relaxation characteristics of three different LOV2 variants, each of which contained a single point mutation that changed a hydrophobic Jα residue (V529) to Ala, Glu or Asn. This residue lies on the hydrophobic surface of the Jα helix proximal to LOV213,15, and we anticipated that each of these changes would perturb the binding equilibrium and thus the relaxation dispersion behavior of the protein. Overall, the 15N-1H HSQC spectra of the mutants were very similar to those of the wild type, suggesting that the mutations had little effect on the overall structure of the protein (data not shown). In addition, backbone 15N CPMG experiments showed that the mutants are dynamic in similar regions as in the wild type protein. However, the mutations have clearly altered the dynamic properties of the system, since the magnitude of Rex for individual resonances varies substantially among them (Fig. 3a). For all resonances, Rex values increased from wild type to V529A to V529E to V529N. Consistent with the data collected for the wild type protein, we were able to globally fit all of the dispersion data for the three mutants to a model for two-site exchange. Figure 3b shows the fitted absolute values of 15N Δωfor all dynamic residues in all constructs. In the majority of cases, we obtained very similar Δω values for a given residue across all four constructs, suggesting that the wild type and mutated proteins all share the same low and high-energy conformations in the dark. Consistent with the rank of Rex magnitudes, the proteins showed increasing populations of the high-energy conformations, from 1.7 ± 0.02% in WT to 2.0 ± 0.03% in V529A to 2.5 ± 0.03% in V529E and 7.6 ± 0.1% in V529N. These observations are consistent with the model that the observed conformational exchange dynamics reflects a binding equilibrium between LOV2 and Jα. Mutations that destabilized the LOV-Jα interaction more significantly than the V529N mutant were difficult to analyze with CPMG-type experiments due to significant line broadening that resulted from more abundant high-energy conformation such that the dynamic residues are no longer observable in the NMR spectrum. For example, only four dynamic residues are observable in CPMG spectra obtained of the Q479T mutant and fits of the corresponding relaxation dispersion curves yield a pB of 22.2%, consistent with the expected increase in the high-energy state population. Analyses of these proteins are also complicated by generally poorer solution behavior than the corresponding wildtype LOV2-Jα construct.
To gain insight into the structure of the high-energy conformation, we used solution NMR spectroscopy to examine the structural properties of a peptide corresponding to an isolated Jα helix and its surrounding elements (AsPhot1 residues 511–545). In the full LOV2-Jα helix construct (AsPhot1 residues 404–560) protein, 13Cα chemical shift values for residues in the Jα helix region are all positive (Fig. 4a), consistent with the α-helical structure indicated by NOEs and backbone 3JHN-Hα coupling constants13. In contrast, the same 13Cα CSI values for the peptide are very close to zero (Fig. 4a), suggesting that Jα exists as a random coil in the absence of stabilizing interactions with the LOV2. To facilitate comparisons of structure of the Jα helix in the LOV2-Jα context and in isolation, we calculated the difference in 15N chemical shifts between the assigned values for Jα peptide and LOV2-Jα protein. This parameter, which we refer to as Δωunfolding, represents the chemical shift change upon unfolding of Jα and dissociation from the LOV2 surface. A comparison of Δωunfolding with ΔωRex, the chemical shift difference between the low and high-energy conformations of the LOV2-Jα protein obtained from the relaxation dispersion measurements above, shows a very high degree of correlation (Fig. 4b). A linear fit of the data gives a slope of 0.57, an intercept of 1.0 and an R-value of 0.8. One possible reason for the fair quality of this fit is that the free Jα peptide and the dark high-energy conformation has different sequence context. In the dark high-energy conformation, the Jα residues are still connected to the LOV2 domain and the extra 20 residues in the C terminus. To address the latter of these concerns, we generated a longer Jα peptide (residue 517–560 vs. 511–545 as used above) that extended to the same C-terminus as the LOV2-Jα construct. Using 15N chemical shift values from this peptide to calculate Δωunfolding of Jα residues, we obtained a linear fit with the same parameters as the shorter peptide.
Given this linear correlation between ΔωRex and Δωunfolding for residues in the Jα helix, our data suggest that the high-energy conformation of LOV2-Jα resembles that of the isolated and unstructured Jαpeptide. That is, since,
Since the 15N chemical shifts of the high-energy conformation approximate those of the Jα peptide, we concluded that the excited state being sampled in the dark has the Jα helix being dissociated from the LOV2 domain and largely unfolded. Thus, the dynamic equilibrium in the dark state involves transient excursions to a higher energy conformation (or conformational ensemble) that closely resembles the photoactivated state of the system.
Our data also indicate that the correlation between ΔωRex and Δωunfolding is not ideal, suggesting that a few Jα residues might retain some low degree of residual structure in the high-energy dark state that is not present in the unfolded peptides, perhaps by maintaining some loose contacts with the LOV core. This would give rise to deviations between ΔωRex and Δωunfolding, as we observed for several residues located at either end of the Jα helix (D522, A523; A542) or in a cluster near its middle (M530, K533, K534) (Fig. 4b). Notably, most of these sites show ΔωRex < Δωunfolding, as is expected if a small degree of residual structure is present in the high-energy dark state. We emphasize that these deviations from linearity are small (<1 ppm 15N) and are not inconsistent with our overall finding that the high-energy dark state has a chiefly-unfolded Jα helix.
Previous analyses of NMR chemical shift dispersion, proteolytic sensitivity and other solution biophysical parameters have suggested that Jα is dissociated from the LOV domain and melted in the lit state13,17,21. Consistent with these observations, 15N-1H HSQC spectra of the isolated Jα peptide and LOV2-Jα protein in the lit state are very similar (Fig. 5a, b). However, the resonance overlap is not exact, and most of the lit state Jα resonances are displaced from their peptide equivalents. A detailed comparison of the dark state, lit state and free Jα peptide spectra reveals that the displacement is not random. Rather, for 13 of the 15 Jα peaks that can be assigned in the lit state spectrum, the lit state position is systematically displaced approximately along a line that connects the corresponding peptide and dark state peaks (Fig. 5c and Fig. 5d; Supplemental Fig. S1). This behavior suggests that in the lit state Jα may be sampling a rapid equilibrium between two conformations, one that resembles the peptide, unfolded and dissociated from the LOV domain, and a second that resembles the predominant dark state structure, folded into an α-helix and bound to LOV2. Such an equilibrium in the lit state would be analogous to that sampled in the dark state, but should be oppositely biased to favor the helix-dissociated conformation. To estimate the populations of the two conformations in the lit state, we assumed that the peptide chemical shifts represent the fully dissociated conformation, and that the dark state chemical shifts of the Jα resonances are identical to those of the helix-bound lit state. While the latter assumption is obviously not exactly correct, the similar behavior of the large majority of Jα resonances indicates it is reasonable. Based on the chemical shifts of the endpoint conformations, the positions of the Jα resonances yield an approximate population of the helix-dissociated conformation of 91 ± 5.7 % in the lit state.
In principle, we should be able to also apply relaxation dispersion analyses to the lit state protein as well to obtain conformational exchange information. Unfortunately, the narrow linewidths of Jα peaks from lit state spectra suggest that these residues are in fast exchange conditions where CPMG methods cannot provide accurate population measurements. Further, lit state samples are only moderately well-behaved over time at high concentration, making it difficult to extract quantitative information from CPMG experiments which require > 48 h data acquisition. Given these issues, we have opted to use the simpler, but more robust, method for measuring population changes by chemical shift changes to estimate the relative pA and pB populations in the lit state.
We have shown that in both the dark and lit states, the LOV2-Jα core of AsPhot1 exists in equilibrium between two major conformations (or more precisely, conformational ensembles): one with Jα bound to LOV2, the other with Jα dissociated away from the rest of the domain. Both conformations are sampled in dark state and illumination state, but to different degrees. Thus, the essential nature of the photoswitch is to change the relative stabilities of these two conformations. In the dark, the helix-bound form is more stable, while this reverses in the light as the helix-dissociated form becomes the more stable form. Relaxation dispersion analyses indicate that the dissociated conformation is populated to only 1.6% in the dark state. Chemical shift analyses of the lit state indicate that FMN adduct formation produces a 91% population of this conformation. Thus, the population of the active state increases ~57-fold upon photo-excitation. Viewed in terms of equilibrium constants (and thus free energy), photo-excitation changes the bound-dissociated Jα equilibrium from 98.4:1.6 in the dark state to 9:91 in the lit state. This corresponds to an overall shift of 620-fold in the bound: dissociated ratio, representing a 3.8 kcal mol−1 change in free energy. We note that this value is more than fifteen-fold smaller than the energy of the blue light photons being absorbed by the internally-bound flavin chromophore, which is ~64 kcal mol−1. Thus, only a small fraction of the total energy of excitation is transmitted to conformational changes in the protein in the form of useful work, while the majority must be dissipated in formation of the flavin-cysteine adduct and as heat.
To confirm these results, we carried out similar measurements on LOV2-Jα protein at 20°C with independently-generated samples. In the dark state, the Jα unbound conformation decreased from 1.6% at 25°C to 0.97% at 20°C, corresponding to 2.7 kcal mol−1 of free energy. In the lit state, the Jα unbound conformation is ~92%, equivalent to ~−1.4 kcal mol−1 free energy. Therefore there are about 4.1 kcal mol−1 of free energy available at 20°C, quite similar to the ~3.8 kcal mol−1 value measured at 25°C. An additional line of independent validation of this value is provided by potential of mean force (PMF) calculations on LOV2-Jα, which indicate a displacement of the Jα helix and a ~3.0 kcal mol−1 change in free energy upon illumination (P. Freddolino, S.M. Harper, K.H.G., K. Schulten; manuscript in preparation).
Previous biochemical and NMR data indicate that dissociation of Jα from LOV2 is sufficient for activation of the AsPhot1 kinase domain18. These observations suggest that in the full length protein the conformational equilibrium in the LOV2-Jα core is coupled to a regulatory equilibrium in the kinase domain. This model suggests that the light-induced shift in the former equilibrium drives activation of the kinase, although the physical mechanisms of this coupling are not currently known. It is difficult in the absence of experimental data to predict how thermodynamic coupling to the rest of the phototropin protein will affect the equilibrium in the LOV2-Jα core, since the covalent connection and interdomain contacts can perturb this equilibrium in unknown ways.
Similarly, the amount by which the equilibrium in the core affects the activity of the kinase domain, in both the dark and lit states, will depend on the specifics of the physical connections. In principle, the actual perturbation of the kinase equilibrium by light could be relatively small, since full activation is likely driven by autophosphorylation, as in other kinases22. The Pak kinase, which is regulated by the Rho GTPases Cdc42 and Rac, appears to be regulated in such a fashion23. GTPase binding to Pak does not substantially disassemble the autoinhibitory apparatus; rather, it simply perturbs the regulatory equilibrium sufficiently to permit autophosphorylation. This covalent modification then fully relieves autoinhibition, raising Pak to a GTPase-independent activation state. On the other hand, there are cases where activity is dictated by the conformational equilibrium. In nitrogen regulator protein C (NtrC), transcription activity of the protein is turned on by phosphorylation of residue D54, which is located in the N-terminal receiver domain. NMR studies on the NtrC receiver domain suggest the existence of an equilibrium between active and inactive conformations in both the phosphorylated and unphosphorylated form24. Mutations that are partially active in the absence of phosphorylation show a close correlation between their activity and population distribution of the equilibrium. Comparison of the population distribution in the phosphorylated versus unphosphorylated form in NtrC in the context of the full length protein will provide insights into the energetic coupling of those “two-component” signaling systems. Similarly, comparison of the LOV2-Jα core and full AsPhot1 in both the dark and lit states will ultimately be necessary to understand how the energy inherent in the photoswitch is used to drive kinase activation.
The ability to reversibly control the activity of arbitrary proteins with light would have tremendous utility in basic and applied research and technology. Although reversible regulation has been achieved in several cases through covalent attachment of small molecule photoswitches25,26, and natural light-controlled ion channels and adenyl cyclases have shown great promise in neuroscience27,28, it would be particularly powerful to generate genetically-encoded photoswitches in a general fashion. Such control might be achieved with chimeras or simple fusions between the LOV2-Jα element of AsPhot1 and various functional elements – e.g. enzymes or binding domains. The success of such an effort is predicated on efficiently coupling the light-dependent changes in free energy provided by a photoswitchable element with other regulatory equilibria. This is analogous to the LOV2 control of kinase activity in phototropin, but such a coupling would have to be achieved through artificial engineering rather than through the course of natural evolution. Lim and colleagues have used an analogous approach to generate synthetic multi-domain proteins with activities controlled by ligand binding29. By fusing a protein and its peptide ligand to an allosterically regulated functional domain, the domain activity can be controlled by free ligand. However, significant suppression of activity (a necessary prerequisite to ligand-mediated activation) was only observed when the intermolecular KD between the protein and ligand was ~10 µM or less, corresponding to 6.8 kcal mol−1 or higher free energy without any covalent linkage between the ligand and the enzyme29. Given that the dark state suppressive free energy of the LOV2-Jα system corresponding to the helix-bound:helix-free conformation is only 2.4 kcal mol−1 with Jα tethered to the LOV domain, more efficient coupling, perhaps achieved through a combination of engineering and screening, will likely be needed to generate LOV2-based photoswitches with a large dynamic range.
Analogous to the LOV2-Jα element of phototropins, photoactive yellow protein (PYP), a blue light photoreceptor from halophilic bacteria, also adopts a PAS fold30. PYP does not bind FMN, but rather uses a covalently linked p-coumaric acid as chromophore. Analogous to Jα in phototropin, PYP has a N-terminal extension that consists of two short α-helices. Numerous studies have shown that photoexcitation leads to unfolding of this N terminal extension31,32. This partially unfolded state is presumed to be the signaling state of PYP. Despite these similarities, relaxation dispersion experiments have not shown evidence of significant µs-ms timescale dynamics in PYP under physiologic conditions (data not shown). Thus, if there are analogous fluctuations to an excited state resembling the lit state in PYP, these are either on timescales outside the 0.1~10 ms range sampled by the CPMG-based technique or the excited state is populated to < ~0.5%, based on the detection limit of the method. Although LOV2-Jα and PYP may utilize similar structurally similar photoswitch mechanisms, the energetics involved in the two systems appear to be distinct. It remains to be discovered whether the structural and energetic properties of the photosensory core of AsPhot1 are conserved among other members of the LOV-containing photoswitch family.
Mutagenesis was performed using the QuikChange site-directed mutagenesis kit (Stratagene). The LOV2-Jα construct consisted of residues 404–560 of Avena sativa phototropin 1. Proteins were expressed from plasmid pHis6-GB113 in E. coli strain BL21(DE3) grown in M9 minimal media containing 1 g/L 15NH4Cl as the sole nitrogen source for uniform 15N labeling and 1 g/L 3-13C- pyruvate33,34 for selective 13C labeling of the following methyl groups: Ala β; Ile γ2, α 1; Leu δ1, δ2; Val γ1, γ2. Expression and purification of all constructs were performed as described13 except for an additional Source 15Q (GE Healthcare) anion exchange chromatography step after an initial Ni2+ affinity chromatography purification. Purified proteins were exchanged into 50 mM phosphate buffer (pH 6), 0.1 M NaCl. All NMR samples contained ~1 mM protein, 10% D2O, 1 mM DTT, 2 mM FMN and a cocktail of protease inhibitors that included EDTA, benzamidine, leupeptine and antipain.
To generate the Jα peptide, AsPhot1 residues G511-E545 were cloned into a modified pET19b vector (Novagen) containing a N-terminal His10-ubiquitin tag and a TEV protease recognition site before the multiple cloning site. The expressed fusion protein was purified using Ni2+-NTA (QIAGEN) affinity followed by anion exchange chromatography (Source 15Q, GE Healthcare). The His-ubiquitin tag was cleaved off by TEV protease and separated from the peptide by Ni2+-NTA affinity chromatography. A Superdex75 (GE Healthcare) gel filtration column was used to buffer exchange the peptide into the same NMR buffer as used for the LOV2-Jα proteins except no FMN was added. Final concentration of the peptide was ~1 mM.
All NMR experiments were performed at 25°C on Varian Inova 600 and 800 MHz spectrometers. Relaxation compensated constant time CPMG pulse sequences were used for both 15N and 13C experiments35–37 using a 40 ms constant time period. All NMR data were processed using NMRDraw38 and NMRview39 software. For each CPMG series, repeats of one or two data points were performed and the intensity of the repeats were used to calculate standard deviation of the series. In cases where the standard deviation was less then 2%, 2% was used in data fitting. For data points with repetition, the average of the two repeats was used for data fitting. Data were fit using software kindly provided by Dr. Lewis Kay, using the following equations20
Where and are the intrinsic R2 values of A and B states.
Dispersion curves were first individually fit assuming the two-site exchange model. Individual fits give similar exchange rates and excited state populations, justifying the use of a two-state global fit, using the same kex and pB for all dynamic residues. Chemical shift values of LOV2-Jα were taken from previous assignments13. Backbone resonance assignments of the peptide were carried out using HNN, HN(C)N41, HNCACB40,42 and CBCA(CO)NH43 experiments. Lit state assignments were obtained as described previously13 using 15N/1H SCOTCH experiments44 that provide 2D correlations of 15Ndark-1Hlit to couple fully assigned 15N/1H HSQC spectra in the dark and the corresponding lit state spectra. This approach provided unambiguous assignments for 12 of the 15 residues in the Jα helix; the other 3 residues that we could assign had ambiguities in the lit state that were resolved by proximity to crosspeaks from spectra acquired on the free Jα peptide.
Assuming the Jα bound state chemical shift values are similar between dark and lit states, the population distribution across the exchange process in the lit state was estimated using backbone 15N and 1H chemical shifts of Jα residues according to the following the equation45:
where A is the Jα bound state in dark, B is the Jα dissociated and unfolded state that is equivalent to the Jα peptide. The chemical shifts of Jα bound state in the dark (state A) is back calculated from the dark state apparent chemical shift values and experimentally determined 98.4% Jα-bound state in the dark using equation (10).
Research was supported by grants from the Welch Foundation to M.K.R. (I–1544) and K.H.G. (I-1424), as well as a grant from the Texas Higher Education Coordinating Board Advanced Technology Program (010019-0124-2003). We thank Dr. Lewis Kay for providing the pulse sequences and software for data acquisition and analyses, Dr. Dmitry M. Korzhnev for help with experimental setup and data fitting, Dr. Dorothee Kern for assistance with improved data acquisition methods. We also thank all members of the Rosen and Gardner labs for helpful discussions.