Although cytochrome P450cam from Pseudomonas putida, the archetype for all heme monooxygenases, has long been known to have a closed active site, recent reports show that the enzyme can also be crystallized in at least two clusters of open conformations. This suggests that the enzyme may undergo significant conformational changes during substrate binding and catalytic turnover. However, these conformations were observed in the crystalline state, and information is needed about the conformations that are populated in solution. In this study, double electron–electron resonance experiments were performed to observe substrate-induced changes in distance as measured by the dipolar coupling between spin labels introduced onto the surface of the enzyme on opposite sides of the substrate access channel. The double electron–electron resonance data show a decrease of 0.8 nm in the distance between spin labels placed at S48C and S190C upon binding the substrate camphor. A rotamer distribution model based on the crystal structures adequately describes the observed distance distributions. These results demonstrate conclusively that, in the physiologically relevant solution state, the substrate-free enzyme exists in the open P450cam-O conformation and that camphor binding results in conversion to the closed P450cam-C form. This approach should be useful for investigating many other P450s, including mammalian forms, in which the role of conformational change is of central importance but not well understood.
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Conformational change and flexibility play important roles in the function of many enzymes. The details of how an enzyme recognizes its substrate and activates its conversion to product are often discussed in terms of lock-and-key, induced fit, or conformational selection models (1–3). Such issues have been of central importance in understanding the substrate specificity of the large family of cytochromes P450 (4, 5). With nearly 12,000 members across all living organisms (6), these heme-containing monooxygenases use an external electron source to activate O2 and insert oxygen into C-H bonds of a wide range of substrates (7). Although P450s exhibit a common protein fold and are believed to use a common reactive Compound I intermediate, they differ vastly in their substrate specificity. For example, many P450s involved in biosynthetic pathways carry out highly regio- and stereo-specific reactions with substrate (8), while other forms, for example those involved in hepatic drug metabolism, can be very indiscriminate by allowing a single isozyme to oxidize a large variety of compounds (9). Much of this diversity is believed to arise from variations in the F and G helices, the intervening loop, and the adjacent B’ helix, which fold around the substrate binding site at the distal heme face. Structurally distinct substrate binding sites have been observed for bacterial (4, 10, 11), microsomal (12–15), and mitochondrial P450s (16). In addition, several P450s have been observed to undergo significant conformational change in these structural elements upon substrate or inhibitor binding (17–22). However, it remains unclear whether conformational change plays an important role in all P450s. Thus, the extent to which substrate recognition is coupled to conformational gating of important functional events including electron transfer and O-O bond cleavage has remained incompletely understood.
Our understanding of the effects of substrate on the conformation of P450cam (CYP101A1), the camphor metabolizing P450 from Pseudomonas putida, has changed over time. Important early studies have suggested that it has an active site that is relatively static in response to substrate binding (4, 10, 23). Based on the comparison of X-ray structures obtained for the enzyme when it is crystallized in the presence of camphor and when small ligands such as dithiothreitol (DTT) are soaked out of the crystals, it was suggested that the enzyme undergoes only minor conformational changes upon binding substrate (10, 23). However, the question of how camphor enters the buried, solvent-inaccessible binding site of the closed form remained unresolved (4). Crystal structures of an open conformation, observed in the presence of large tethered substrate analogs, provided a proposal for the substrate access route (24, 25) through a substrate channel, but it remained initially uncertain how much these tethered substrate analogs were perturbing the native conformation. Thus, it was significant when P450cam was reported to visit the same open conformation when crystallized in the absence of any substrate (26). This suggested that crystal packing interactions may have prevented substrate-dependent conformational changes in the experiments where DTT was soaked out of crystals (23). A recent study of the structures of P450cam bound to a family of tethered substrates showed that the enzyme can be observed in at least three distinct clusters of conformations, representing open (P450cam-O), intermediate (P450cam-I), and closed (P450cam-C) states (27) that result primarily from segmental movements of the F and G helices. These observations have provided support for the proposal that in solution, P450cam undergoes significant conformational changes in the F and G helices during substrate binding and turnover. However, these conformations were observed for the protein in the crystalline state, and information is currently lacking about the conformations that are actually populated in the physiologically relevant solution state. This issue is particularly important given the recent report of NMR studies (28), which demonstrate significant changes in the conformational ensemble upon camphor binding, but did not reveal large movements in the F and G helices. Therefore, it has become critically important to establish whether the previously reported open conformations are seen in the solution state.
In this report, double electron–electron resonance (DEER) spectroscopy of spin-labeled P450cam is used to clarify these issues by providing conformational information for P450cam in solution, specifically determining the position of the F-G loop relative to the rest of the enzyme in the presence and absence of camphor. This study also demonstrates how this method may prove generally useful for probing conformational changes in many other forms of P450 for which such information is both desirable and difficult to obtain.
Results
Two spin labels were introduced onto the surface of P450cam to allow observation of the change from the open to closed conformation in solution by measurement of the inter-spin through-space magnetic dipolar coupling. To facilitate the use of thiol-specific spin labels, surface accessible wild-type cysteines were first mutated to serine. Wild-type P450cam contains eight cysteine residues. C334, the most reactive surface cysteine, is routinely changed to alanine for studies by our laboratory and many others to prevent intermolecular disulfide formation (29). Four cysteines (C58, C85, C136, and C285) that are at least partially exposed on the surface of the enzyme were also replaced with serine to give P450cam-(4S) containing no surface accessible thiols. The remaining three cysteines are not reactive: C148 and C242 are partially buried, and C357 is coordinated to the heme. A comparison of the crystal structures for camphor-bound P450cam-C and camphor-free P450cam-O (Fig. 1) shows that the distance between the F-G loop and the residues across the substrate channel (around helix A and strand β1) is maximally sensitive to changes in the open-to-closed transition. S48 and S190 are surface-exposed serines with their side chains directed toward solvent and were thus considered good candidates for mutation to cysteine and spin label attachment. S48, near the A helix, does not change position relative to the core of the protein during the conformational change, while residue S190, in the middle of the F-G loop, moves toward S48 by 0.8 nm upon closure of the substrate channel (Fig. 1). Thus, the S48C and S190C mutations were introduced into the P450cam-(4S) background to generate P450cam-(4S,2C). Samples of purified P450cam-(4S) and P450cam-(4S,2C) protein displayed 0.01 and 1.82 reactive thiols, respectively, as shown by the Ellman reaction (30). UV-visible spectra showed that the ferric heme in P450cam-(4S, 2C) is converted from a low-spin state (λmax = 417 nm) in the absence of camphor to a predominantly high-spin state (λmax = 392 nm) upon addition of 1 mM camphor (Fig. S1), consistent with the characteristic behavior of wild-type P450cam. Following spin labeling of P450cam-(4S,2C) with the perdeuterated spin label MTSL-d15 (d15-2,5-dihydro-2,2,5,5-tetramethyl-3-[[(methylsulfonyl)thio]methyl]-1H-pyrrol-1-yloxy) and removal of unreacted spin label by gel filtration, approximately 1.6 spin labels per protein were observed by double integration of CW EPR spectra (Fig. S2) and comparison to a CuEDTA standard. Finally, the crystal structure at 2.0-Å resolution was determined for spin-labeled P450cam-(4S,2C) in the camphor-bound closed form (Table S1). The electron density for one of two molecules in the asymmetric unit clearly indicated the presence of the spin labels extending from the S48C and S190C sulfur atoms. This density fit well to a single conformation of MTSL built into the model, from which the inter-spin distance in the closed form was measured at 4.4 nm (Fig. 2). Electron density for the spin labels in the second molecule of the asymmetric unit were not as clearly defined suggesting some conformational disorder.
Fig. 1.
Crystal structures of P450cam in the closed substrate-bound (cyan, pdb 2CPP) and open substrate-free (orange, PDB ID code 3L61) conformations. For clarity, only the most displaced regions, including the helices F and G and the connecting loop, are colored. Cα positions are labeled at the S48C and S190C mutation sites. Heme is shown in red.
Fig. 2.
Crystal structure of spin-labeled P450cam-(4S,2C) in the closed substrate-bound conformation. (A) Spin labeled mutant (magenta, PDB ID code 4EK1) and wild type (cyan, PDB ID code 2CPP) are superimposed. For clarity, only secondary structures around the spin labeling sites and the I helix are shown. (B) 2Fo-Fc map of the labeling sites, contoured at 1σ.
DEER experiments were performed to measure substrate-induced changes in magnetic dipolar coupling between the two spin labels on P450cam. Fig. 3A shows the experimental DEER time traces. The frequencies of the oscillations are determined by the magnetic dipolar coupling between the unpaired electrons of the two spin labels which, in turn, is proportional to the inverse cube of the distance rNO between the centers of the spin label N-O bonds, νdip = (52 MHz)(rNO/nm)-3. The higher frequency for the substrate-bound form clearly demonstrates a reduced distance between the two spin labels compared to the substrate-free form. In both cases, the significant damping of the echo modulation amplitude over time indicates that there is a heterogeneous distribution of distances rather than a single distance. The corresponding distance distributions, extracted from the time traces using Tikhonov regularization (31), are shown in Fig. 3B and are dominated by peaks at 5.6 nm and 4.8 nm, respectively, for samples in the absence and presence of camphor. The distance distributions therefore show a 0.8-nm reduction in distance upon substrate binding. This is in agreement with the difference predicted from the crystal structures for conversion from the P450cam-O to the P450cam-C conformation (Fig. 1). Smaller peaks in the distance distribution at approximately 3 nm in Fig. 3B may result from artifacts due to deuterium modulation of the spin labels, or from effects of the coupling between the spin labels and the Fe(III) center. The small peak at approximately 8 nm for the substrate-free form is assigned to incomplete removal of inter-molecular modulation background. For the closed conformation, the distance of 4.8 nm observed experimentally by DEER (Fig. 3B) is in reasonable agreement with the value of 4.4 nm observed in the crystal structure of the labeled protein (Fig. 2).
Fig. 3.
(A) Experimental (color) and fitted (black) DEER time traces for substrate-free (−cam, blue) and camphor-bound (+cam, red) cytochrome P450cam-(4S,2C) spin labeled at positions S48C and S190C. (B) DEER distance distributions as a function of the distance between the NO groups of the two spin labels (rNO), obtained from Tikhonov regularization of the experimental DEER traces.
Modeling of the spin label side-chain conformations and their disorder on the surface of the enzyme largely accounts for the distance distribution width, providing additional insight. Starting from the crystal structures of open and closed states, the rNO distributions were estimated computationally using a conformational distribution model based on a rotameric library for MTSL (32) and the crystal structures of the unlabeled enzyme, which were kept rigid. Fig. 4A illustrates the predicted disorder of the NO spatial location at the S48C and S190C labeling sites, resulting in the distance distributions shown by dotted lines in Fig. 4B and summarized in Tables S2–S4. The modeled distributions of Fig. 4B remain 0.3–0.4 nm shorter than the experimentally determined DEER distance distributions. In the camphor free state, the width of the observed distribution is very similar to that modeled by conformational flexibility of the 5 dihedral angles within MTSL (32). In the camphor bound state, the observed distribution is a little narrower than that modeled. Although these effects might be sensitive to the glass-transition temperature of the solvation shell around the protein, assumed to be 175 K in the calculations (33), or to partial orientation selection in the experiment (34), these results suggest that the distribution widths of rNO observed in the DEER experiments are dominated by torsional disorder of the spin label rather than by the protein backbone Cα–Cα distance distribution.
Fig. 4.
(A) Rotameric models of substrate-free (−cam, PDB ID code 3L61) and substrate-bound (+cam, PDB ID code 2CPP) forms of cytochrome P450cam spin-labeled with MTSL at positions S48C and S190C. The spheres indicate the positions of the NO group in the various rotamers, and sphere diameters are proportional to the fractional populations. (B) Comparison of modeled (dashed) and experimental (solid) distance distributions for substrate-bound and substrate-free P450cam. The gray line represents the observed distance between NO bond centers in the crystal structure of spin-labeled P450cam +cam (PDB 4EK1; see Fig. 2).
Discussion
Role of Conformational Change.
Despite much effort over many decades, the molecular basis of substrate specificity in P450s is not fully understood, and recent results suggest a shifting paradigm that may have broad impact on our understanding of their function. Even P450cam, the most well studied example, has recently been trapped in multiple conformations in response to binding of tethered substrate analogues (27). These results suggest P450cam might be similar to other P450 enzymes, which have been observed to undergo significant conformational changes in response to substrate or ligand binding (35–38). Still, it remains possible that the substrate channel of some P450s is more flexible than others and thus, the role played by conformational change and dynamics in defining substrate specificity of a particular form remains unresolved. The current study clarifies these issues for the solution state of P450cam and suggests that DEER spectroscopy may be used with this and other forms of P450 to gain a greater insight into these questions.
Crystal vs. Solution.
The recent observation of multiple conformations of P450cam in response to binding a library of tethered substrates (27) demonstrates the range of structures that can be sampled by a given form of the enzyme and suggests how substrates enter and leave the binding site of the closed conformation. However, these states have been characterized in the crystalline state, and additional information about which conformations are populated in solution is needed. The different structures obtained for the substrate-free enzyme depending on whether the substrate is soaked out of crystals (23) or is absent during crystallization (26) suggest that crystal contacts might influence the conformational equilibrium and warn that the states seen in the crystal do not necessarily reflect those populated in solution. In addition, while a recently reported NMR study (28) has shown that camphor binding results in alteration of the conformational ensemble in solution, large-scale movements of the F and G helices were not indicated. It was suggested that crystal contacts may be responsible for this difference, but it is possible that the presence of CO in the NMR studies may also contribute to changing the conformational equilibrium. A complete picture will likely require a combination of techniques. Thus, the use of DEER spectroscopy to quantitatively observe conformational changes in solution provides crucial new information about the basic function of this model enzyme.
Distances and Their Changes.
For the camphor bound closed conformation, the absolute distance between spin labels determined by DEER is in good agreement with the crystal structure of the spin-labeled protein. In general, the inter-spin distances are about 0.5–0.7 nm longer than the backbone distances between the residues (see Table S3) due to the tether of the two spin label side chains. By themselves, the absolute distance values are therefore difficult to interpret. On the other hand, the clear change in spin label distance observed by DEER in response to substrate binding is direct evidence for substrate-induced closure of the substrate channel. The similarity of the substrate-induced -0.8 nm distance change observed by DEER and that predicted from X-ray crystallography strongly supports the conclusion that the substrate-free enzyme exists in the open P450cam-O conformation and that camphor binding results in conversion to the closed P450cam-C form. Thus, in the present case, the conformational states characterized by X-ray crystallography appear to accurately reflect the forms populated in solution, provided that the crystals are grown in equilibrium with the final substrate concentration. The DEER data show that the substrate-free and substrate-bound forms in solution are predominantly in the open and closed conformations, respectively (Fig. 3B). In addition, the DEER data provide no evidence for a significant population of the P450cam-I intermediate conformation observed in the presence of some tethered substrates (27).
Distance Distributions.
An important feature of DEER spectroscopy is its ability to provide information about the distribution of distances in the sample. A heterogeneity in the distance results in the damping of the echo modulation amplitude, which can be characterized if the acquired data extends long enough in time. In the present case, careful optimization of experimental conditions, high purity of the samples and the use of deuterated buffer components and spin labels (see Methods) allowed DEER data collection up to 5 μs, enough to observe significantly more than one full cycle of echo modulation for both samples. As a result, distance distributions were obtained in which not only the distribution modes but also the distribution widths are significant. However, the DEER traces are too short to allow interpretation of the shape of the distance distributions. Further studies on deuterated protein (39) may allow refinement of DEER measurements such that better resolved distribution shapes are obtained and substrate binding events can be probed beyond the simple two-state conformational model used here. This could possibly yield evidence for the multi-step conformational pathway that has recently been proposed for this enzyme (27).
Rotamer Analysis.
As shown in Fig. 4B and in SI Text, the rotameric modeling based on the crystal structures yielded distance distributions that are in reasonable agreement with experiment. The distances predicted based on the crystal structures of the unlabeled enzyme are slightly shorter than those experimentally observed by DEER in the solution state. Remarkably, when the crystal structure of the spin-labeled enzyme was used for distance distribution modeling, the predicted and experimental distances distribution modes coincide (Fig. S3), suggesting that small changes in the local structure around the labeling sites can affect the accuracy of rotamer modeling. The systematic deviation of the predicted distance distributions from the experimental DEER results might be a result of the approximate nature of the rotamer library approach, which is known to yield biased results in some cases (32, 40). Also, it is possible that the use of the crystal structures in place of the unknown solution structures contributes to the discrepancy because the regions around one or both labeling sites might have solution conformations that are different from those in the crystal structure (Figs. S4 and S5). On the other hand, the spin label side chain might interact with specific residues on the protein surface in a way not captured by the rotamer model (e.g., via hydrogen bonds), and thus only a subset of the modeled rotameric states shown in Fig. 4 might be populated. In the case of a specific label-protein interaction, some of the width of the observed distance distributions could then arise from heterogeneity of protein backbone conformation. This possibility is under investigation and can be tested by additional DEER studies on mutants near the labeling sites.
Mechanism.
Based on our results it seems very likely that substrate binding in P450cam is facilitated by the conformational flexibility of the enzyme. In the absence of substrate, the open form P450cam-O appears to be energetically most favored. Based on present data, it is not clear whether this is the only conformation that is (statically) populated at physiological temperature or whether there is dynamic exchange within an ensemble of thermally accessible conformations. The latter would be consistent with the DEER data only if the conformations do not differ significantly in their 48–190 distances. Upon passage of the substrate through the channel into the active site, the energetics of the conformational landscape is changed such that the closed form P450cam-C is most stable. This open-closed transition can either be a substrate-directed induced-fit process or a dynamic relaxation of the enzyme into the new conformational energetic landscape. Structural studies using tethered substrate analogues (27) suggest a prominent intermediate conformation state along the open-closed transition, with a possible role of putidaredoxin as conformational or functional effector (41–43). Conformational changes associated with substrate recognition, enzyme reduction and O2 binding may be coupled to the position of the catalytically important water and Thr-252 side chain (44). It has been shown that the product complex closely resembles the closed substrate-bound conformation (45), and thus product release would necessitate reopening of the substrate channel. The release mechanism and its trigger are currently not fully understood.
In conclusion, the DEER-based coarse-grain structural analysis approach used in this work provides new insight into the mechanism of substrate binding in P450cam. It should be useful for investigating conformational states of many other P450s, for example mammalian forms, that are proposed to undergo conformational changes during substrate recognition, but for which key structural information is lacking.
Methods
Mutant Construction and Protein Purification.
Mutations were introduced into the plasmid DNA (pETCAM-C334A) for Escherichia coli expression of P450cam using methods previously described and verified by DNA sequencing. In preparation for site-specific spin-labeling of P450cam, reactive surface exposed cysteines were first removed by mutation to serine. These sites included C58S, C85S, C136S, and C285S in addition to the C334A mutation normally used by our laboratory to prevent protein dimerization. The resulting construct, labeled P450cam-(4S), was used as a background to introduce a pair of surface cysteines at residues S48 and S190 to generate the construct denoted P450cam-(4S, 2C) for spin label attachment.
Protein expression and purification utilized methods previously described for P450cam-C334A (26). One mM dithiothreitol (DTT) was included in all buffers to prevent intermolecular disulfide bond formation. Protein stock solutions were stored at -80 °C in 50 mM potassium phosphate (pH 6.0), 300 mM KCl, 1 mM camphor and 1 mM DTT. The Ellman reaction was used to determine the number of reactive surface thiols on the protein. Briefly, protein was incubated (5 min at RT) in 100 μM DTNB (5,5′-dithio-bis(2-nitrobenzoic acid)), 1 mM camphor and 100 mM Tris (pH 8.0). The optical absorbance at 412 nm (ε412 = 13,600 M-1 cm-1) was used for calculation of concentration of free thiol concentration and was corrected for the contribution of heme to the absorbance using control samples omitting DNTB.
Spin Labeling and EPR Sample Preparation.
P450cam-(4S, 2C) was labeled with either MTSL (2,5-dihydro-2,2,5,5-tetramethyl-3-[[(methylsulfonyl)thio]methyl]-1H-pyrrol-1-yloxy, (1-oxyl-2,2,5,5-tetramethyl-Δ3-pyrroline-3-methyl) methanethiosulfonate) (Toronto Research Chemicals) or its perdeuterated derivative, MTSL-d15. DTT was first removed from the protein by buffer exchange into 50 mM Tris (pH 7.4) by gel filtration using two sequential PD-10 columns (GE Healthcare). Camphor (1 mM) and KCl (200 mM) were added before incubating for 10 min at 4 °C in a 10-fold molar excess of the spin label. The reaction was quenched by removal of excess of MTSL by gel filtration through a PD-10 column. EPR samples of spin-labeled ferric P450cam were prepared at a final protein concentration of ca. 150 μM by exchange into buffer made from 99% D2O containing 50 mM Tris (pD 7.8), 200 mM KCl, 1 mM camphor (omitted in the substrate-free sample) and 30% d8-glycerol (Cambridge Isotopes Laboratory). For each sample, 150 μL was loaded into a 4 mm o.d. quartz tube (Wilmad 706-PQ-9.50) and flash frozen in liquid nitrogen.
Crystallization, Data Collection, and Structure Determination.
Crystals of spin-labeled P450cam-(4S,2C) were grown from 50 mM Tris, pH 7.4, 200 mM KCl, 1 mM camphor and 12% PEG 8000. Crystals were transferred to the mother liquor with additional 25% PEG 600 and flash frozen on nylon loops. X-ray diffraction data were collected at 100 K on the beamline 7-1 at the Stanford Synchrotron Radiation Laboratory. Data processing, molecular replacement, refinement, and validation were carried out with the same methods employed previously (26). Statistics for data collection and refinement are shown in Table S1. The atomic coordinates and structure factors have been deposited in the Protein Data Bank (PDB ID code 4EK1). Two protein molecules are present in the asymmetric unit. MTSL was modeled into observed electron density maps at S48C and S190C in molecule A and at S48C in molecule B but not at S190C of molecule B due to disorder. Structural illustrations were generated using the Pymol Molecular Graphics System (46).
EPR Data Collection and Analysis.
X-band continuous-wave (CW) EPR spectra were measured at the CalEPR Center at UC Davis on a Bruker ECS106 spectrometer at 9.5 GHz/0.34 T using a Bruker SHQE resonator and an Oxford liquid helium cooling system. The EPR signals of low- and high-spin ferric iron from the heme (47) did not interfere with the nitroxide measurements because the Fe3+ signals in the nitroxide region are very weak due to their large anisotropy compared to the nitroxide signal. The concentration of spin labels was determined by comparison of the double integral of the spin label spectrum to the double integral of a CuEDTA solution with known concentration.
X-band DEER data were acquired at the CalEPR Center at UC Davis on a Bruker EleXsys E580 spectrometer at 9.5 GHz using a Bruker MD4 dielectric resonator and an Oxford liquid helium cooling system, at temperatures between 20 and 50 K and a magnetic field of about 340 mT. The DEER pulse sequence (π/2)1 - τ1 - (π)1 - τ1 + t - (π)2 - (τ2 - t) - (π)1 - τ2 - [echo] was used with pulse lengths of 16/32/32 ns for the probe pulses (subscript 1) and 32 ns for the pump pulse (subscript 2). The value of τ1 was adjusted to a maximum in the 2H nuclear modulation envelope (400 ns), and t was varied between -40 ns and about 5 μs in increments of 12 or 20 ns. Averaging over τ1, which could cancel potential distance artifacts due to deuterium or proton modulations (48), could not be performed due to an insufficient signal-to-noise ratio. The pump frequency ν2 was centered in the resonator mode and aligned with the spectral maximum. The probe frequency ν1 was 65 MHz above ν2.
Deuteration of the buffer was crucial for the DEER measurements because it prolonged the phase memory time, TM, allowing data acquisition in which t extended to 5 μs (total pulse sequence length almost 11 μs). Additional perdeuteration of the spin label gave an additional 10% increase in TM, but also introduced deep modulations as a function of τ1.
For the substrate-bound form, the best data were obtained at 50 K. At lower temperatures, the modulations were shallower and the damping was stronger, probably due to saturation effects. For the substrate-free form, relaxation at 50 K was too fast to allow the acquisition of an entire 5 μs long time trace, so data were acquired at 30 K where relaxation is slower.
EPR spectral simulations and quantitations were performed using EasySpin 4.0 (49). The DEER data were analyzed and fit with DeerAnalysis2011 (31). Rotameric modeling was performed with MMM2011 (32).
Data Availability
Data deposition: The atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.pdb.org (PDB ID code 4EK1).
ACKNOWLEDGMENTS.
The authors acknowledge the assistance of J. Rosenberg and helpful discussions with Professors T.L. Poulos, J.R. Halpert, E.F. Johnson, C.D. Stout, and J.B. Ames. This work was supported by the National Institutes of Health (GM41049 to D.B.G.) and the Department of Energy (DE-FG02-09ER16117 to R.D.B.). Portions of this research were carried out at the Stanford Synchrotron Radiation Lightsource, a Directorate of SLAC National Accelerator Laboratory and an Office of Science User Facility operated for the U.S. Department of Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research, and by the National Institutes of Health, National Institute of General Medical Sciences (including P41GM103393) and the National Center for Research Resources (P41RR001209). The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of NIGMS, NCRR or NIH.
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The authors acknowledge the assistance of J. Rosenberg and helpful discussions with Professors T.L. Poulos, J.R. Halpert, E.F. Johnson, C.D. Stout, and J.B. Ames. This work was supported by the National Institutes of Health (GM41049 to D.B.G.) and the Department of Energy (DE-FG02-09ER16117 to R.D.B.). Portions of this research were carried out at the Stanford Synchrotron Radiation Lightsource, a Directorate of SLAC National Accelerator Laboratory and an Office of Science User Facility operated for the U.S. Department of Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research, and by the National Institutes of Health, National Institute of General Medical Sciences (including P41GM103393) and the National Center for Research Resources (P41RR001209). The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of NIGMS, NCRR or NIH.
Author contributions: D.B.G., R.D.B., Y.-T.L., and S.S. designed the research; S.S., Y.-T.L., and M.Z. performed the research; Y.-T.L. and R.F.W. provided new materials and sample characterization; S.S. analyzed the data; and S.S., Y.-T.L., D.B.G. and R.D.B. wrote the paper.
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S.S. and Y.-T.L. contributed equally to this work.
Competing Interests
The authors declare no conflict of interest.
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S. Stoll, Y. Lee, M. Zhang, R.F. Wilson, R.D. Britt, D.B. Goodin, Double electron–electron resonance shows cytochrome P450cam undergoes a conformational change in solution upon binding substrate, Proc. Natl. Acad. Sci. U.S.A.
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