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19F NMR reveals the conformational properties of free thrombin and its zymogen precursor prethrombin-2

Open AccessPublished:May 01, 2020DOI:https://doi.org/10.1074/jbc.RA120.013419
      The conformational properties of trypsin-like proteases and their zymogen forms remain controversial because of a lack of sufficient information on their free forms. Specifically, it is unclear whether the free protease is zymogen-like and shifts to its mature form upon a ligand-induced fit or exists in multiple conformations in equilibrium from which the ligand selects the optimal fit via conformational selection. Here we report the results of 19F NMR measurements that reveal the conformational properties of a protease and its zymogen precursor in the free form. Using the trypsin-like, clotting protease thrombin as a relevant model system, we show that its conformation is quite different from that of its direct zymogen precursor prethrombin-2 and more similar to that of its fully active Na+-bound form. The results cast doubts on recent hypotheses that free thrombin is zymogen-like and transitions to protease-like forms upon ligand binding. Rather, they validate the scenario emerged from previous findings of X-ray crystallography and rapid kinetics supporting a pre-existing equilibrium between open (E) and closed (E*) forms of the active site. In this scenario, prethrombin-2 is more dynamic and exists predominantly in the E* form, whereas thrombin is more rigid and exists predominantly in the E form. Ligand binding to thrombin takes place exclusively in the E form without significant changes in the overall conformation. In summary, these results disclose the structural architecture of the free forms of thrombin and prethrombin-2, consistent with an E*–E equilibrium and providing no evidence that free thrombin is zymogen-like.

      Introduction

      The trypsin fold defines the structural architecture of a mature “active” protease and its immature “inactive” zymogen precursor and has been studied in considerable detail, both functionally and structurally (
      • Page M.J.
      • Di Cera E.
      Serine peptidases: classification, structure and function.
      ,
      • Hedstrom L.
      Serine protease mechanism and specificity.
      ,
      • Perona J.J.
      • Craik C.S.
      Structural basis of substrate specificity in the serine proteases.
      ). The zymogen-to-protease transition is described by the Huber–Bode mechanism (
      • Huber R.
      • Bode W.
      Structural basis of the activation and action of trypsin.
      ) and involves a proteolytic cleavage at a conserved Arg residue in the so-called activation domain, followed by insertion of the new N terminus into the protein core and folding of the active site. The mechanism was originally assumed to generate a fully functional protease from its inactive zymogen precursor, but this view proved too simplistic when structures of trypsin-like proteases started to accumulate in the Protein Data Bank (PDB).
      The trypsin fold has long been assumed to be mostly “rigid” (
      • Bode W.
      • Fehlhammer H.
      • Huber R.
      Crystal structure of bovine trypsinogen at 1–8 A resolution: I. Data collection, application of patterson search techniques and preliminary structural interpretation.
      ,
      • Bode W.
      • Huber R.
      Crystal structure analysis and refinement of two variants of trigonal trypsinogen: trigonal trypsin and PEG (polyethylene glycol) trypsinogen and their comparison with orthorhombic trypsin and trigonal trypsinogen.
      ,
      • Bode W.
      • Schwager P.
      • Huber R.
      The transition of bovine trypsinogen to a trypsin-like state upon strong ligand binding: the refined crystal structures of the bovine trypsinogen-pancreatic trypsin inhibitor complex and of its ternary complex with Ile-Val at 1.9 A resolution.
      ,
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ). However, the entire “west wall” of the active site defined by the amino acid segment at positions 215–217 adopts different conformations that open and close access to the primary specificity pocket at the bottom of the active-site region. This conformational plasticity is of functional significance (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ). In the open conformation (E), the active site enables ligand binding with high affinity. In the closed conformation (E*), no binding is possible. The active site is occluded by collapse of the side chain of Trp215 and a shift of the backbone of the segment at positions 215–217. The aperture leading to the primary specificity pocket and defined by the Cα–Cα distance of residues Gly193 and Gly216 changes from 12 Å in the E form to only 8.1 Å in the E* form (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ). In the D216G mutant of αI-tryptase (
      • Rohr K.B.
      • Selwood T.
      • Marquardt U.
      • Huber R.
      • Schechter N.M.
      • Bode W.
      • Than M.E.
      X-ray structures of free and leupeptin-complexed human αI-tryptase mutants: indication for an α → β-tryptase transition.
      ), chymotrypsinogen (
      • Wang D.
      • Bode W.
      • Huber R.
      Bovine chymotrypsinogen A X-ray crystal structure analysis and refinement of a new crystal form at 1.8 A resolution.
      ), and the thrombin precursor prethrombin-2 (
      • Pozzi N.
      • Chen Z.
      • Zapata F.
      • Pelc L.A.
      • Barranco-Medina S.
      • Di Cera E.
      Crystal structures of prethrombin-2 reveal alternative conformations under identical solution conditions and the mechanism of zymogen activation.
      ), both the E and E* forms are detected in the same crystal. Independent support of alternative conformations for the free form of protease and zymogen comes from rapid kinetics studies of ligand binding to the active site. The mechanism of recognition for small tripeptides is consistent with a pre-existing E*–E equilibrium of conformational selection rather than induced fit (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ). The E* form prevails in the zymogen and gradually shifts to the E form during transition to the mature protease (
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ). In fact, the E*–E equilibrium complements the Huber–Bode mechanism and contributes to organization of the active site for efficient binding and catalysis (
      • Stojanovski B.M.
      • Chen Z.
      • Koester S.K.
      • Pelc L.A.
      • Di Cera E.
      Role of the I16-D194 ionic interaction in the trypsin fold.
      ).
      Although evidence of the E*–E equilibrium from the current structural database (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Pozzi N.
      • Vogt A.D.
      • Gohara D.W.
      • Di Cera E.
      Conformational selection in trypsin-like proteases.
      ) and rapid kinetics (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ,
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ) is solid, direct proof that such conformational plasticity exists in solution remains elusive. Structural features revealed by X-ray may be biased by crystal packing or by the extreme solution conditions often necessary to achieve crystallization. Kinetic experiments provide unequivocal evidence of conformational selection only when certain conditions are met for the observed relaxation rates (
      • Vogt A.D.
      • Di Cera E.
      Conformational selection or induced fit?: A critical appraisal of the kinetic mechanism.
      ). Previous NMR studies on the clotting protease thrombin in different bound states (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ) have identified a progressive rigidification of the enzyme upon ligation. From these studies, Huntington (
      • Huntington J.A.
      Slow thrombin is zymogen-like.
      ) and Krishnaswamy and co-workers (
      • Kamath P.
      • Huntington J.A.
      • Krishnaswamy S.
      Ligand binding shuttles thrombin along a continuum of zymogen-like and proteinase-like states.
      ) have concluded that thrombin is inherently plastic and shuttles within an ensemble of conformations that are disordered and zymogen-like when free but rigid and protease-like when bound (
      • Huntington J.A.
      Slow thrombin is zymogen-like.
      ,
      • Kamath P.
      • Huntington J.A.
      • Krishnaswamy S.
      Ligand binding shuttles thrombin along a continuum of zymogen-like and proteinase-like states.
      ). However, this ensemble view of thrombin (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ,
      • Huntington J.A.
      Slow thrombin is zymogen-like.
      ,
      • Kamath P.
      • Huntington J.A.
      • Krishnaswamy S.
      Ligand binding shuttles thrombin along a continuum of zymogen-like and proteinase-like states.
      ) remains a speculation largely inconsistent with existing X-ray (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Pozzi N.
      • Vogt A.D.
      • Gohara D.W.
      • Di Cera E.
      Conformational selection in trypsin-like proteases.
      ) and rapid kinetics (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ,
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ) data and lacks validation from NMR studies of free thrombin or of the zymogen precursor prethrombin-2. Whether free thrombin is zymogen-like and switches to the mature conformation by induced fit or pre-exists in alternative conformations from which the ligand selects the optimal fit can only be established by studies of the free form. Recent NMR measurements by the Komives group (
      • Handley L.D.
      • Fuglestad B.
      • Stearns K.
      • Tonelli M.
      • Fenwick R.B.
      • Markwick P.R.
      • Komives E.A.
      NMR reveals a dynamic allosteric pathway in thrombin.
      ) have targeted the “apo-form” of thrombin using the S195M mutant and compared the dynamics with those of thrombin bound at the active site (
      • Fuglestad B.
      • Gasper P.M.
      • Tonelli M.
      • McCammon J.A.
      • Markwick P.R.
      • Komives E.A.
      The dynamic structure of thrombin in solution.
      ). However, these studies have been carried out in the presence of Na+ and therefore describe the dynamics of ligation of the fully active Na+-bound form of thrombin rather than its free form.
      In this study, we investigate for the first time the conformation of the protease thrombin and its zymogen precursor prethrombin-2 in the free form using 19F NMR. We labeled all nine Trp residues in thrombin and prethrombin-2 to interrogate the conformational properties of the protein in the absence of any added ligand and Na+. 19F NMR has been used for a number of biologically important systems (
      • Bann J.G.
      • Pinkner J.
      • Hultgren S.J.
      • Frieden C.
      Real-time and equilibrium 19F-NMR studies reveal the role of domain-domain interactions in the folding of the chaperone PapD.
      ,
      • Danielson M.A.
      • Falke J.J.
      Use of 19F NMR to probe protein structure and conformational changes.
      ,
      • Frieden C.
      • Hoeltzli S.D.
      • Bann J.G.
      The preparation of 19F-labeled proteins for NMR studies.
      ,
      • Manglik A.
      • Kim T.H.
      • Masureel M.
      • Altenbach C.
      • Yang Z.
      • Hilger D.
      • Lerch M.T.
      • Kobilka T.S.
      • Thian F.S.
      • Hubbell W.L.
      • Prosser R.S.
      • Kobilka B.K.
      Structural insights into the dynamic process of β2-adrenergic receptor signaling.
      ,
      • Aramini J.M.
      • Hamilton K.
      • Ma L.C.
      • Swapna G.V.T.
      • Leonard P.G.
      • Ladbury J.E.
      • Krug R.M.
      • Montelione G.T.
      19F NMR reveals multiple conformations at the dimer interface of the nonstructural protein 1 effector domain from influenza A virus.
      ) and is ideally suited for studying the Trp residues of thrombin that are known to report on changes linked to the E*–E equilibrium, the zymogen-to-protease conversion, and ligand binding (
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ,
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ,
      • Bah A.
      • Garvey L.C.
      • Ge J.
      • Di Cera E.
      Rapid kinetics of Na+ binding to thrombin.
      ).

      Results

      19F labeling

      We labeled all Trp residues of thrombin and prethrombin-2 at the 5 position of the indole ring with 19F and solved the X-ray structures at 2.3 and 2.1 Å resolution, respectively (Table 1). The nine Trp residues are distributed over the entire surface and function as effective reporters of the conformational state of the protein. The structures show all Trp residues correctly labeled without significant perturbation of the overall architecture (Fig. 1). Extra density detected at the 5 position of the indole ring in all cases supports uniform labeling of the reagents used for NMR studies (Fig. 2). The 19F-labeled prethrombin-2 structure features a conformation of the active site similar to that of WT (
      • Pozzi N.
      • Chen Z.
      • Zapata F.
      • Pelc L.A.
      • Barranco-Medina S.
      • Di Cera E.
      Crystal structures of prethrombin-2 reveal alternative conformations under identical solution conditions and the mechanism of zymogen activation.
      ) (RMSD = 0.33 Å). The 19F-labeled thrombin structure bound to the active site inhibitor PPACK and Na+ is very similar to the unlabeled complex (
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ,
      • Pineda A.O.
      • Carrell C.J.
      • Bush L.A.
      • Prasad S.
      • Caccia S.
      • Chen Z.W.
      • Mathews F.S.
      • Di Cera E.
      Molecular dissection of Na+ binding to thrombin.
      ) (RMSD = 0.41 Å).
      Table 1Crystallographic data for 19F-labeled prethrombin-2 and PPACK-inhibited thrombin
      PDB entry6V5T6V64
       Buffer/salt0.1 m HEPES, pH 7.00.2 m sodium/potassium tartrate, pH 7.5
       PEG8000 (25%)3350 (14%)
      Data collection
       Wavelength (Å)1.541.54
       Space groupP21P21212
       Unit cell dimensions (Å)a = 44.5, b = 58.9, c = 52.4, β = 98.4a = 61.9, b = 86.6, c = 50.5
       Molecules/asymmetric unit11
       Resolution range (Å)40–2.140–2.3
       Observations79,52162,626
       Unique observations15,69612,020
       Completeness (%)99.3 (97.0)94.9 (84.5)
      Rsym (%)7.3 (55.9)11.5 (33.4)
      I/σ(I)18.0 (2.4)11.7 (2.4)
      Refinement
       Resolution (Å)40–2.140–2.3
      Rcryst, Rfree0.177, 0.2300.197, 0.277
       Reflections (working/test)14,911/77211,333/588
       Protein atoms2,3562,283
       Solvent molecules103108
       PPACK1
       Na+2
       RMSD bond lengths (Å)
      RMSD from ideal bond lengths and angles and RMSD in B-factors of bonded atoms.
      0.0080.010
       RMSD angles (°)
      RMSD from ideal bond lengths and angles and RMSD in B-factors of bonded atoms.
      1.51.8
       RMSD ΔB (Å2) (mm/ms/ss)
      mm, main chain–main chain; ms, main chain–side chain; ss, side chain–side chain.
      3.21/2.98/3.502.04/2.20/2.08
       Protein41.543.6
       Solvent42.640.6
       PPACK32.3
       Na+33.8
       Ramachandran plot (%)
      Most favored95.095.0
      Generously allowed5.05.0
      Disallowed0.00.0
      a RMSD from ideal bond lengths and angles and RMSD in B-factors of bonded atoms.
      b mm, main chain–main chain; ms, main chain–side chain; ss, side chain–side chain.
      Figure thumbnail gr1
      Figure 1Crystal structures of 19F-labeled prethrombin-2 (A) and PPACK (cyan sticks) inhibited thrombin in the Na+-bound (purple ball) form (B) with the side chains of the nine Trp residues of the protein shown as sticks. Residues Trp51 and Trp215 feature characteristic NMR resonances and dynamics (see Figure 4, Figure 5) and are indicated, along with residues Trp141, Trp148, and Trp60d. The 19F label on the 5 position of the indole ring is clearly visible for all Trp residues (see also ). The two structures are similar to unlabeled prethrombin-2 (
      • Pozzi N.
      • Chen Z.
      • Zapata F.
      • Pelc L.A.
      • Barranco-Medina S.
      • Di Cera E.
      Crystal structures of prethrombin-2 reveal alternative conformations under identical solution conditions and the mechanism of zymogen activation.
      ) (RMSD = 0.33 Å) and PPACK-bound thrombin (
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ,
      • Pineda A.O.
      • Carrell C.J.
      • Bush L.A.
      • Prasad S.
      • Caccia S.
      • Chen Z.W.
      • Mathews F.S.
      • Di Cera E.
      Molecular dissection of Na+ binding to thrombin.
      ) (RMSD = 0.41 Å), proving that labeling introduced no bias in the fold. Details of the structures are given in .
      Figure thumbnail gr2
      Figure 2Details of the 19F-labeled Trp residues of prethrombin-2 (A) and thrombin (B), taken from the crystal structures shown in (see also ). Extra density on the 5 position of the indole side chain is clearly detected for all nine Trp residues, demonstrating that labeling was uniform. The electron density 2FoFc map (green mesh) is contoured at 1 σ.

      19F NMR measurements

      Having established uniform 19F labeling by X-ray crystallography, we proceeded to collect 1D 19F NMR spectra for thrombin and prethrombin-2 in the free form, devoid of ligands bound to the active site or Na+. These conditions have not been explored in previous studies (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ,
      • Handley L.D.
      • Fuglestad B.
      • Stearns K.
      • Tonelli M.
      • Fenwick R.B.
      • Markwick P.R.
      • Komives E.A.
      NMR reveals a dynamic allosteric pathway in thrombin.
      ,
      • Fuglestad B.
      • Gasper P.M.
      • Tonelli M.
      • McCammon J.A.
      • Markwick P.R.
      • Komives E.A.
      The dynamic structure of thrombin in solution.
      ) but are essential to determine the intrinsic properties of the protein in the free form. Overall, the 1D 19F spectra show seven well-dispersed peaks for thrombin and only four for prethrombin-2 (Fig. 3A and Table 2). The most striking differences between zymogen and protease are the resonance at −43.5 ppm for thrombin not seen in prethrombin-2, and the range between −47.2 and −49.0 ppm where thrombin shows four distinct peaks, but prethrombin-2 features only two, one large and broad (−47.9 ppm) and the other smaller (−48.6 ppm). A sharp resonance observed in prethrombin-2 around −49.8 ppm is replaced by a smaller one in thrombin, slightly shifted to −49.4 ppm. The presence of well-defined and separable peaks in thrombin as opposed to prethrombin-2 suggests that most of the Trp residues in the zymogen experience a similar chemical environment. However, this conclusion is not supported by the crystal structure (Fig. 1) where some Trp residues are exposed to solvent (Trp60d, Trp148, and Trp215) and others are more buried (Trp51). An alternative explanation is that Trp residues in prethrombin-2 exchange among multiple conformations leading to broad, overlapping linewidths. Hence, thrombin likely explores a smaller conformational space and is intrinsically more rigid than its zymogen precursor prethrombin-2. The observation points out significant differences between protease and zymogen in the free form and does not support recent claims of free thrombin being zymogen-like (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ,
      • Huntington J.A.
      Slow thrombin is zymogen-like.
      ,
      • Kamath P.
      • Huntington J.A.
      • Krishnaswamy S.
      Ligand binding shuttles thrombin along a continuum of zymogen-like and proteinase-like states.
      ,
      • Fuglestad B.
      • Gasper P.M.
      • Tonelli M.
      • McCammon J.A.
      • Markwick P.R.
      • Komives E.A.
      The dynamic structure of thrombin in solution.
      ). In fact, free thrombin is way more similar to its Na+-bound form (Fig. 3B) than its zymogen precursor prethrombin-2 (Fig. 3A). Rapid kinetics studies suggest that a significant fraction of the free enzyme exists in the E form (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ). Structural studies document almost complete overlap between the free and bound E forms (
      • Pineda A.O.
      • Carrell C.J.
      • Bush L.A.
      • Prasad S.
      • Caccia S.
      • Chen Z.W.
      • Mathews F.S.
      • Di Cera E.
      Molecular dissection of Na+ binding to thrombin.
      ,
      • Vogt A.D.
      • Pozzi N.
      • Chen Z.
      • Di Cera E.
      Essential role of conformational selection in ligand binding.
      ). The addition of Na+ is known to boost the catalytic activity of the enzyme (
      • Wells C.M.
      • Di Cera E.
      Thrombin is a Na+-activated enzyme.
      ) and to rigidify the structure (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ,
      • Pozzi N.
      • Chen R.
      • Chen Z.
      • Bah A.
      • Di Cera E.
      Rigidification of the autolysis loop enhances Na+ binding to thrombin.
      ). The 19F NMR spectra in Fig. 3B show that binding of Na+ sharpens and better separates the peaks of free thrombin and removes the peak at −47.9 ppm. We conclude that free thrombin is not zymogen-like. Rather, it is quite distinct from its zymogen precursor prethrombin-2 and already contains features of its more rigid, Na+-bound form as predicted by a mechanism of conformational selection (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Vogt A.D.
      • Di Cera E.
      Conformational selection or induced fit?: A critical appraisal of the kinetic mechanism.
      ,
      • Vogt A.D.
      • Bah A.
      • Di Cera E.
      Evidence of the E*–E equilibrium from rapid kinetics of Na+ binding to activated protein C and factor Xa.
      ,
      • Vogt A.D.
      • Di Cera E.
      Conformational selection is a dominant mechanism of ligand binding.
      ).
      Figure thumbnail gr3
      Figure 3Overlay of 1D NMR spectra between prethrombin-2 and thrombin (A) and between free thrombin and thrombin bound to Na+ (B). The spectra show how free thrombin is more similar to its Na+-bound form than the zymogen precursor prethrombin-2. The difference is particularly noticeable in the number of separate peaks detected for zymogen and protease, consistent with a more rigid structure for the latter.
      Table 219F chemical shifts (ppm) of Trp residue
      ThrombinPrethrombin-2
      Trp29NDND
      Trp51−46.7−46.7
      Trp60d−48.5, −48.7−47.9
      Trp96−47.9−47.9
      Trp141−43.5−47.9
      Trp148−48.5−47.9
      Trp207ND−47.9
      Trp215−47.5−47.9, −49.8
      Trp237−49.4−48.6

      Resonance assignment

      Given the differences between zymogen and protease in the free form, we turned our attention to the specific Trp residues responsible for the observed changes. Assignments of the nine Trp residues in both thrombin and prethrombin-2 (Fig. 4, A and B) were made from spectra for which each individual Trp residue was replaced by Phe. The substitution is inconsequential on the catalytic properties and specificity of thrombin and has been used to identify the fluorophores responsible for Na+ binding to the enzyme (
      • Bah A.
      • Garvey L.C.
      • Ge J.
      • Di Cera E.
      Rapid kinetics of Na+ binding to thrombin.
      ). Assignment of individual Trp residues was often complicated by the lack of selective perturbation of peaks in the WT spectrum. In the case of prethrombin-2, the W215F substitution affects both the large peak around −47.9 ppm and the peak at −49.8 ppm, almost 2 ppm apart (Fig. 5A). This suggests that Trp215 exists in alternative conformations that exchange very slowly. Mutations of Trp60d, Trp96, Trp141, Trp148, Trp207, and Trp215 result in perturbation of the peak at −47.9 ppm (Fig. 4A). Residues Trp51 and Trp237 map to the peaks at −46.7 and −48.6 ppm, respectively, but Trp29 and Trp207 could not be assigned. In the case of thrombin (Fig. 4B), clustering is less pronounced, and residues Trp141, Trp51, Trp215, Trp96, and Trp237 could be assigned to resonances at −43.5, −46.7, −47.5, −47.9, and −49.4 ppm, respectively (see Fig. 5B for W215F). The relative solvent exposure of these residues is consistent with the crystal structure (
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ,
      • Pineda A.O.
      • Chen Z.W.
      • Caccia S.
      • Cantwell A.M.
      • Savvides S.N.
      • Waksman G.
      • Mathews F.S.
      • Di Cera E.
      The anticoagulant thrombin mutant W215A/E217A has a collapsed primary specificity pocket.
      ) (see also Fig. 1B). Trp51 maps to the same resonance position as in prethrombin-2 (Fig. 4A), but Trp237 is shifted upfield. Of the remaining four Trp residues, Trp29 and Trp207 could not be assigned, whereas Trp148 and Trp60d cluster in the peaks within the range of −47 to −48 ppm and could not be separated, suggesting similar solvent exposure as seen in the crystal structure (
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ,
      • Pineda A.O.
      • Chen Z.W.
      • Caccia S.
      • Cantwell A.M.
      • Savvides S.N.
      • Waksman G.
      • Mathews F.S.
      • Di Cera E.
      The anticoagulant thrombin mutant W215A/E217A has a collapsed primary specificity pocket.
      ) (see also Fig. 1B). We conclude that prethrombin-2 is more dynamic than thrombin, with most of its Trp residues in slow exchange between alternative conformations and experiencing an environment that changes significantly during the conversion to thrombin.
      Figure thumbnail gr4
      Figure 41D NMR spectra of prethrombin-2 (A) and thrombin (B) with peaks assigned from single site replacement of Trp with Phe (see also ). The substitution does not change the functional properties and specificity of thrombin (
      • Bah A.
      • Garvey L.C.
      • Ge J.
      • Di Cera E.
      Rapid kinetics of Na+ binding to thrombin.
      ). A, assignment of individual Trp residues in prethrombin-2 often did not result in selective perturbation of peaks in the spectrum (A). The W215F replacement affected both the large peak around −47.9 ppm and the peak at −49.8 ppm (). Mutations of Trp60d, Trp96, Trp141, Trp148, Trp207, and Trp215 perturbed the peak at −47.9 ppm, and those of Trp51 and Trp237 affected the peaks at −46.7 and −48.6 ppm, respectively. Trp29 and Trp207 could not be assigned. B, clusters are less pronounced than in prethrombin-2. Mutations of Trp141, Trp51, Trp215, Trp96, and Trp237 mapped to peaks at −43.5, −46.7, −47.5, −47.9, and −49.4 ppm, respectively. Trp51 mapped to the same resonance position as in prethrombin-2, but Trp237 was shifted upfield. Of the remaining four Trp residues, Trp29 and Trp207 could not be assigned, and Trp148 and Trp60d clustered in the peaks in the range of −47 to −48 ppm and could not be separated.
      Figure thumbnail gr5
      Figure 5Overlay of 1D NMR spectra between WT (blue) and mutant W215F (red) for prethrombin-2 (A) and thrombin (B). Asterisks indicate the region of the spectrum perturbed by the single-site replacement.

      Residue dynamics

      Individual resonances could be assigned to residues Trp51 and Trp215 in thrombin and Trp51 in prethrombin-2. These residues were investigated further by measurements of T1, T2, and CPMG relaxation dispersion to gain insight into their range of motions. Trp51 is positioned 33 Å away from the Na+ binding site and 22 Å away from the catalytic Ser195 (Fig. 1). The peak for Trp51 has the same resonance position at −46.6 ppm in both thrombin and prethrombin-2 (Fig. 4) and broadens from 0.27 to 0.37 Hz relative to thrombin bound to Na+ (Fig. 6), suggesting the presence of multiple conformations. Indeed, the peak shows a distinct relaxation dispersion profile indicative of conformational exchange, especially in prethrombin-2 (Figure 7, Figure 8). Collection at a second field of 600 MHz allowed relaxation dispersion curves at both fields to be fit to a two-state model in the fast-exchange regime (
      • Mazur A.
      • Hammesfahr B.
      • Griesinger C.
      • Lee D.
      • Kollmar M.
      ShereKhan: calculating exchange parameters in relaxation dispersion data from CPMG experiments.
      ) with kex = 19,000 ± 1,000 s−1 in thrombin and kex = 2,970 ± 20 s−1 in prethrombin-2 (Fig. 9). The intrinsic dynamics of Trp51 indicate faster exchange in thrombin than prethrombin-2. Interestingly, the exchange in thrombin is completely abrogated upon Na+ binding (Fig. 8), suggesting rigidification of a residue located 33 Å away. The slower exchange at Trp51 observed in prethrombin-2 is indicative of the presence of more large-scale motions compared with thrombin. These findings add complexity to the scenario emerged from the X-ray structural database where the conformation of Trp51 is essentially the same in prethrombin-2 (
      • Pozzi N.
      • Chen Z.
      • Zapata F.
      • Pelc L.A.
      • Barranco-Medina S.
      • Di Cera E.
      Crystal structures of prethrombin-2 reveal alternative conformations under identical solution conditions and the mechanism of zymogen activation.
      ) and thrombin free or bound to ligands (
      • Pineda A.O.
      • Carrell C.J.
      • Bush L.A.
      • Prasad S.
      • Caccia S.
      • Chen Z.W.
      • Mathews F.S.
      • Di Cera E.
      Molecular dissection of Na+ binding to thrombin.
      ). We conclude that Na+ binding has long-range effects on the structure of thrombin and that Trp51 is allosterically coupled to regions affected by the zymogen to protease conversion, as well as Na+ binding, thereby establishing a new allosteric pathway of communication within the protein that affects widely separated residues.
      Figure thumbnail gr6
      Figure 6Overlay of resonances for Trp51 in prethrombin-2, free thrombin, and thrombin bound to Na+ indicating distinct line broadening of Trp51.
      Figure thumbnail gr7
      Figure 7CPMG relaxation dispersion profiles in the fast time-scale regime of residues Trp51 (filled circles) and Trp215 (open circles) in prethrombin-2 (A) and thrombin (B).
      Figure thumbnail gr8
      Figure 8Field-dependent 19F CPMG relaxation dispersion data for Trp51 in prethrombin-2 (filled circles), free thrombin (open circles), or thrombin bound to Na+ (half-filled circles).
      Figure thumbnail gr9
      Figure 9Field-dependent CPMG relaxation dispersion profiles in the fast time-scale regime. Filled circles represent transverse relaxation rates (R2) as a function of υcpmg acquired at 658.780 MHz, and open circles represent transverse relaxation rates (R2) as a function of υcpmg acquired at 564.686 MHz. The data were fit to a two-state model in the fast-exchange regime yielding the following: A, residue Trp51 in prethrombin-2, kex = 2,970 ± 20 s−1, R20 (564.686 MHz) = 204 ± 1 s−1, R20 (658.780 MHz) = 260 ± 0.3 s−1. B, residue Trp215 in thrombin. kex = 7,980 ± 90 s−1, R20 (564.686 MHz) = 160 ± 1 s−1, R20 (658.780 MHz) = 258 ± 1 s−1. C, residue Trp51 in thrombin, kex = 19,000 ± 1,000 s−1, R20 (564.686 MHz) = 100 ± 10 s−1, R20 (658.780 MHz) = 140 ± 10 s−1.
      Residue Trp215 defines the P3 site of recognition for substrate binding to the active site (
      • Bode W.
      • Turk D.
      • Karshikov A.
      The refined 1.9-A X-ray crystal structure of d-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships.
      ) and exists in different conformations that open and close access to the active site according to the X-ray structural database (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ,
      • Pozzi N.
      • Vogt A.D.
      • Gohara D.W.
      • Di Cera E.
      Conformational selection in trypsin-like proteases.
      ). The role of the indole side chain of Trp215 in the E*–E equilibrium has been tested by rapid kinetics with the W215A mutation and found not to be responsible for the opening and closing of access to the primary specificity pocket (
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ). Specifically, the W215A mutant binds ligands at the active site with a mechanism of conformational selection as WT, proving that removal of the side chain of Trp215 does not equalize access to the active site between the E* and E forms. The role of the side chain of Trp215 is to keep the active site open and slow down the E → E* conversion by establishing an interaction with the benzene ring of Phe227 (
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ). Once this hydrophobic interaction is disrupted, closure of the active site in the E* form is faster and results in reduced catalytic activity (
      • Arosio D.
      • Ayala Y.M.
      • Di Cera E.
      Mutation of W215 compromises thrombin cleavage of fibrinogen, but not of PAR-1 or protein C.
      ,
      • Marino F.
      • Pelc L.A.
      • Vogt A.
      • Gandhi P.S.
      • Di Cera E.
      Engineering thrombin for selective specificity toward protein C and PAR1.
      ). The rate of exchange between E* and E in thrombin and prethrombin-2 becomes of interest. Trp215 maps to a single peak at −47.5 ppm in thrombin (Figs. 4B and 5B). In prethrombin-2, Trp215 maps with other residues to the broad peak in the range of −47 to −48 ppm and also to a unique peak close to −49.8 ppm (Figs. 4A and 5A). The presence of two peaks separated by a large chemical shift indicates that Trp215 features two distinct conformations that exchange very slowly or not at all in prethrombin-2 (Fig. 7). The dynamic profile changes upon transition to thrombin and supports fast exchange between two states (Fig. 7). Measurements at different field strength (Fig. 9) yield a rate of exchange kex = 7,980 ± 90 s−1. Although the structural database documents a similar behavior for Trp215 in zymogen and protease with regard to the E* and E forms controlling access to the primary specificity pocket (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ,
      • Pozzi N.
      • Vogt A.D.
      • Gohara D.W.
      • Di Cera E.
      Conformational selection in trypsin-like proteases.
      ), the dynamics of Trp215 are consistent with an exchange considerably faster in thrombin than prethrombin-2.

      Discussion

      Our understanding of the conformational nature of trypsin-like proteases and their zymogens has been deeply influenced by the celebrated Huber–Bode mechanism of zymogen activation (
      • Huber R.
      • Bode W.
      Structural basis of the activation and action of trypsin.
      ). Activity is assumed to result from proteolytic cleavage of a conserved Arg residue in the activation domain followed by an ionic interaction that is established between the new N terminus of Ile16 and the side chain of the highly conserved Asp194. The newly formed H-bond between Ile16 and Asp194 organizes the oxyanion hole around Gly193 and the catalytic Ser195 and the primary specificity pocket around Asp189. This transition, however, is neither necessary nor sufficient to generate a fully active protease. Activity can be triggered by alternative mechanisms. Single-chain tissue-type plasminogen activator features catalytic activity by establishing an intramolecular H-bond that produces the same structural transitions as the Huber–Bode mechanism (
      • Renatus M.
      • Engh R.A.
      • Stubbs M.T.
      • Huber R.
      • Fischer S.
      • Kohnert U.
      • Bode W.
      Lysine 156 promotes the anomalous proenzyme activity of tPA: X-ray crystal structure of single-chain human tPA.
      ). A similar strategy is used by the plasminogen activator in the saliva of Desmodus rotundus (
      • Renatus M.
      • Bode W.
      • Huber R.
      • Stürzebecher J.
      • Prasa D.
      • Fischer S.
      • Kohnert U.
      • Stubbs M.T.
      Structural mapping of the active site specificity determinants of human tissue-type plasminogen activator: implications for the design of low molecular weight substrates and inhibitors.
      ). Bacteria have evolved proteins like streptokinase (
      • Wakeham N.
      • Terzyan S.
      • Zhai P.
      • Loy J.A.
      • Tang J.
      • Zhang X.C.
      Effects of deletion of streptokinase residues 48–59 on plasminogen activation.
      ) and staphylocoagulase (
      • Friedrich R.
      • Panizzi P.
      • Fuentes-Prior P.
      • Richter K.
      • Verhamme I.
      • Anderson P.J.
      • Kawabata S.
      • Huber R.
      • Bode W.
      • Bock P.E.
      Staphylocoagulase is a prototype for the mechanism of cofactor-induced zymogen activation.
      ) that can activate the host fibrinolytic and coagulation cascades without proteolytic cleavage of their target zymogens plasminogen or prothrombin. An entire class of zymogen activator peptides mimicking streptokinase and staphylocoagulase has been developed by phage display (
      • Landgraf K.E.
      • Steffek M.
      • Quan C.
      • Tom J.
      • Yu C.
      • Santell L.
      • Maun H.R.
      • Eigenbrot C.
      • Lazarus R.A.
      An allosteric switch for pro-HGF/Met signaling using zymogen activator peptides.
      ). The Huber–Bode mechanism also appears not to be sufficient for protease function. A pre-existing equilibrium between closed (E*) and open (E) conformations of the active site controls the onset of substrate binding and catalysis (
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ). In the E* form, substrate cannot bind to the active site, and catalysis is impeded. Importantly, the balance between E* and E changes between zymogen and protease, with the E* form predominating in the former (
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ,
      • Stojanovski B.M.
      • Chen Z.
      • Koester S.K.
      • Pelc L.A.
      • Di Cera E.
      Role of the I16-D194 ionic interaction in the trypsin fold.
      ) and presaging little overlap between the free forms of protease and its zymogen precursor.
      The results reported in this study offer a view of the structural architecture of thrombin and prethrombin-2 in the free form that is entirely consistent with the E*–E equilibrium. We find no evidence that free thrombin is zymogen-like, as speculated in previous functional (
      • Lechtenberg B.C.
      • Johnson D.J.
      • Freund S.M.
      • Huntington J.A.
      NMR resonance assignments of thrombin reveal the conformational and dynamic effects of ligation.
      ,
      • Huntington J.A.
      Slow thrombin is zymogen-like.
      ,
      • Kamath P.
      • Huntington J.A.
      • Krishnaswamy S.
      Ligand binding shuttles thrombin along a continuum of zymogen-like and proteinase-like states.
      ) and computational (
      • Kahler U.
      • Kamenik A.S.
      • Kraml J.
      • Liedl K.R.
      Sodium-induced population shift drives activation of thrombin.
      ) studies. Labeling all nine Trp residues of the protein for 19F NMR measurements show that free thrombin is quite different from prethrombin-2 and more similar to its Na+-bound form, in agreement with the structural differences between E* and E (
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Pineda A.O.
      • Carrell C.J.
      • Bush L.A.
      • Prasad S.
      • Caccia S.
      • Chen Z.W.
      • Mathews F.S.
      • Di Cera E.
      Molecular dissection of Na+ binding to thrombin.
      ), the predominance of E* for prethrombin-2 and of E for thrombin (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Vogt A.D.
      • Di Cera E.
      Conformational selection or induced fit?: A critical appraisal of the kinetic mechanism.
      ), and the fact that E form changes little upon ligand binding (
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Vogt A.D.
      • Pozzi N.
      • Chen Z.
      • Di Cera E.
      Essential role of conformational selection in ligand binding.
      ).
      Our 19F NMR data provide information on the dynamics of critical Trp residues of the protein. Most of these residues cannot be assigned in the 1D 19F NMR spectrum of prethrombin-2 because of overlap of linewidths. Trp51 maps to a single peak and features rapid exchange between alternative conformations in both prethrombin-2 and thrombin. Trp215 maps to two widely separated peaks, indicating a very slow exchange between alternative conformations. When prethrombin-2 transitions to thrombin upon activation, the overall structure becomes less dynamic, with several individual peaks in the 1D 19F NMR spectrum that can be assigned to specific Trp residues. Unlike Trp51, residue Trp215 features distinct dynamics from prethrombin-2 and exchanges rapidly between alternative conformations with kex = 7,980 ± 90 s−1, which is significantly faster than the rate for the E*–E exchange detected by rapid kinetics (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ,
      • Chakraborty P.
      • Acquasaliente L.
      • Pelc L.A.
      • Di Cera E.
      Interplay between conformational selection and zymogen activation.
      ). Although the dynamic nature of Trp215 (Fig. 5B) is consistent with alternative conformations documented by the structural database (
      • Gohara D.W.
      • Di Cera E.
      Allostery in trypsin-like proteases suggests new therapeutic strategies.
      ,
      • Niu W.
      • Chen Z.
      • Gandhi P.S.
      • Vogt A.D.
      • Pozzi N.
      • Pelc L.A.
      • Zapata F.
      • Di Cera E.
      Crystallographic and kinetic evidence of allostery in a trypsin-like protease.
      ,
      • Pozzi N.
      • Vogt A.D.
      • Gohara D.W.
      • Di Cera E.
      Conformational selection in trypsin-like proteases.
      ), its fast time scale of exchange points to events that eventually do not influence access to the active site as documented in the E*–E equilibrium. Indeed, removal of the indole of Trp215 with the W215A substitution changes little the E*–E distribution compared with WT thrombin (
      • Pelc L.A.
      • Koester S.K.
      • Chen Z.
      • Gistover N.E.
      • Di Cera E.
      Residues W215, E217 and E192 control the allosteric E*–E equilibrium of thrombin.
      ,
      • Chakraborty P.
      • Di Cera E.
      Induced fit is a special case of conformational selection.
      ). Other features of the protein, like movement of the backbone of the 215–217 segment, may be responsible for the E*–E equilibrium (
      • Vogt A.D.
      • Chakraborty P.
      • Di Cera E.
      Kinetic dissection of the pre-existing conformational equilibrium in the trypsin fold.
      ) and should be investigated further by NMR of the free form to extend the work reported in this study.

      Materials and methods

      Reagents

      Prethrombin-2 cloned into a pet28 Escherichia coli expression vector was transformed into the BL21DE3 E. coli strain. 50-ml starter cultures in LB supplemented with ampicillin were grown for up to 16 h at 37 °C in an orbital shaker rotating at 225 rpm. Starter cultures were diluted 1:50 in LB, also supplemented with ampicillin, and grown for a further 3–4 h until A600 > 1.0 was reached. The growth culture was spun down at 4,000 rpm for 20 min. The pellet was then dissolved in minimal medium for incorporation of 5-F-Trp and grown for an additional 2 h before recombinant protein expression was induced by adding 1 mm isopropyl β-d-thiogalactopyranoside and growing at 25 °C overnight. The formulation for 5-F-Trp minimal media reads as 50 mm Na2HPO4, 25 mm KH2PO4, 20 mm NH4Cl, 100 μg/ml ampicillin, 0.25 mg/liter 5-F-Trp, and 0.4% w/v d-glucose.
      For inclusion body purification and refolding, the cells were pelleted by centrifugation at 4,000 rpm for 25 min and resuspended in 50 mm Tris, pH 7.4, 20 mm EDTA, 1 mm DTT, and 1% Triton X-100. The cells were lysed using an Avestin C3 emulsified or sonicator. Inclusion bodies were separated by centrifugation at 10,000 rpm for 15 min. Inclusion bodies were washed sequentially in 50 mm Tris, pH 7.4, 20 mm EDTA, and 1 m NaCl followed by 50 mm Tris, pH 7.4, and 20 mm EDTA. Inclusion bodies were solubilized in 40 ml of 7 m guanidine HCl and homogenized using a Dounce homogenizer. Inclusion bodies were centrifuged at 10,000 rpm for 10 min to remove insoluble material. Inclusion bodies were refolded by dilution into 50× excess of 50 mm Tris, 500 mm NaCl, 1 mm EDTA, 10% glycerol, 600 mm Arg, 0.2% Brij58, 1 mml-Cys, pH 8.3. Inclusion bodies were incubated at 25 °C overnight.
      Properly folded prethrombin-2 was purified using a heparin-affinity column and buffer-exchanged through a size-exclusion column or 10-kDa Centricon into a final NMR buffer formulation of 20 mm Tris, 700 mm NaCl, or 700 mm choline chloride, 50 mm Arg, and 10% trehalose. Thrombin was generated from prethrombin-2 using ecarin and exchanged into the same NMR buffer. Protein was concentrated to ∼5 mg/ml and spiked with 10% D2O for 19F NMR data collection.
      All constructs for NMR data collection contained neutralization of the active-site Ser195 with the S195A mutation to prevent autocatalytic degradation and enable collection of relaxation dispersion data over a period of days.

      19F NMR studies

      1D 19F NMR measurements were carried out at 658.780 MHz, 25 °C using a 19F QCI cryoprobe (City University of New York Advanced Science Research Center Biomolecular NMR Facility) and a 5-mm PFG quadruple resonance inverse detection cryoprobe (Saint Louis University High Resolution NMR Facility). No differences with 1D 19F NMR spectra were observed between the probes. Relaxation dispersion NMR experiments were carried out at 658.780 MHz, 25 °C (City University of New York Advanced Science Research Center Biomolecular NMR Facility) with a 19F QCI cryoprobe and at a second field strength of 564.686 MHz with a TCI (H/F-CN-D) cryogenic probe (Wisconsin). All spectra were referenced to TFA. Typical 1D 19F acquisition parameters were a 20,000-Hz sweep width (42.5 ppm), 0.35-s acquisition time, 5-s relaxation delay time, and 5.0-μs 90° pulse length. Spectra were processed with a 20-Hz exponential line broadening using topspin.
      Longitudinal (T1) and transverse (T2) 19F relaxation measurements were determined using classic 1D inversion recovery (T1) and the Carr–Purcell–Meiboom–Gill (CPMG) spin echo pulse sequence (T2) (
      • Carr H.Y.
      • Purcell E.M.
      Effects of diffusion on free precession in nuclear magnetic resonance experiments.
      ,
      • Meiboom S.
      • Gill D.
      Modified spin-echo method for measuring nuclear relaxation times.
      ). 19F T1 inversion recovery experiments were acquired at a series of variable delay times (0.0625, 0.125, 0.25, 0.5, 1.0, 2.0, 4.0, and 8.0 s) and a relaxation delay of 7 s. 19F CPMG experiments consisted of a 90x − [τcp − 180y − τcp] n pulse train acquired with a series of spin-echo evolution times (e.g. 8 points ranging between 0.5 and 128 ms). Longitudinal and transverse relaxation times were computed by fitting plots of 19F signal intensity for a given 5-F-Trp residue or peak as a function of variable decay (T1) or spin-echo times (T2). 19F CPMG relaxation dispersion experiments in which the transverse relaxation rate, R2, is determined as the function of the delay between 180° pulses (2τcp) were acquired across a series of τcp values (e.g. 50–500 μs) and plotted against CPMG frequency (
      • Aramini J.M.
      • Hamilton K.
      • Ma L.C.
      • Swapna G.V.T.
      • Leonard P.G.
      • Ladbury J.E.
      • Krug R.M.
      • Montelione G.T.
      19F NMR reveals multiple conformations at the dimer interface of the nonstructural protein 1 effector domain from influenza A virus.
      ). All measurements were collected in duplicate.

      X-ray studies

      Crystallization of 19F-labeled, WT prethrombin-2, and thrombin bound to the active site inhibitor H-d-Phe-Pro-Arg-CH2Cl (PPACK) was achieved at 25 °C by the vapor diffusion technique, using an Art Robbins Instruments PhoenixTM liquid handing robot with 6–10 mg/ml protein (0.3 μl) mixed with an equal volume reservoir solution. Optimization of crystal growth was achieved by the hanging-drop vapor-diffusion method mixing 3 μl of protein with equal volumes of reservoir solution (see Table 1). The crystals were grown in 1 week at 25 °C and frozen with 25% glycerol from the original mother liquor. X-ray diffraction data were collected at 100 K with a home source (Rigaku 1.2 kw MMX007 generator with VHF optics) Rigaku Raxis IV2+ detector and were indexed, integrated, and scaled with the HKL2000 software package (
      • Otwinowski Z.
      • Minor W.
      Processing of x-ray diffraction data collected by oscillation methods.
      ). Structures were solved by molecular replacement using PHASER from the CCP4 suite (
      • Dodson E.J.
      • Winn M.
      • Ralph A.
      Collaborative Computational Project, number 4: providing programs for protein crystallography.
      ) and the structures of human prethrombin-2 mutant S195A (PDB code 3SQH) and human thrombin in complex with PPACK (PDB code 1PPB) as starting models. Refinement and electron density generation were performed with REFMAC5 from the CCP4 suite. 5% of the reflections were randomly selected as a test set for cross-validation. Model building and analysis were carried out using COOT (
      • Emsley P.
      • Cowtan K.
      Coot: model-building tools for molecular graphics.
      ). In the final refinement stage, TLS tensors modeling rigid-body anisotropic temperature factors were calculated and applied to the model for the 19F-labeled thrombin bound to PPACK. Ramachandran plots were calculated using PROCHECK (
      • Morris A.L.
      • MacArthur M.W.
      • Hutchinson E.G.
      • Thornton J.M.
      Stereochemical quality of protein structure coordinates.
      ). The statistics for data collection and refinement are summarized in Table 1.

      Data availability

      Atomic coordinates and structure factors for the two structures reported in the manuscript have been deposited in the PDB (accession code 6V5T for 19F-labeled prethrombin-2 and accession code 6V64 for 19F-labeled thrombin bound to PPACK). All other data described in the manuscript are contained within the manuscript.

      Acknowledgments

      An earlier investigation of thrombin and prethrombin-2 by 19F NMR was carried out in 2008 and 2009 by Prafull S. Gandhi as part of his Ph.D. thesis at Washington University in St. Louis. The data reported in this study were collected at the National Magnetic Resonance Facility in Madison, the City University of New York Advanced Science Research Center Biomolecular NMR Facility; and the Saint Louis University High Resolution NMR Facility. The National Magnetic Resonance Facility at Madison is supported by NIH grant P41GM103399 (NIGMS), old number: P41RR002301, with equipment purchased with funds from the University of Wisconsin-Madison, the NIH (P41GM103399, S10RR02781, S10RR08438, S10RR023438, S10RR025062, S10RR029220), the NSF (DMB-8415048, OIA-9977486, BIR-9214394), and the USDA. We are grateful to Tracey Baird for help with illustrations.

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