BBAGEN-28106; No of Pages 8; 4C: 2, 3, 5, 6 Biochimica et Biophysica Acta xxx (2014) xxx–xxx

Contents lists available at ScienceDirect

Biochimica et Biophysica Acta journal homepage: www.elsevier.com/locate/bbagen

Review

Real-time protein NMR spectroscopy and investigation of assisted protein folding☆ Amit Kumar, Jochen Balbach ⁎ Institut für Physik, Biophysik, und Mitteldeutsches Zentrum für Struktur und Dynamik der Proteine (MZP), Martin-Luther Universität Halle-Wittenberg, Halle D-06120, Germany

a r t i c l e

i n f o

Article history: Received 26 September 2014 Received in revised form 26 November 2014 Accepted 2 December 2014 Available online xxxx Keywords: Real-time NMR Protein folding Kinetic intermediates Molecular chaperones

a b s t r a c t Background: During protein-folding reactions toward the native structure, short-lived intermediate states can be populated. Such intermediates expose hydrophobic patches and can self-associate leading to non-productive protein misfolding. A major focus of current research is the characterization of short-lived intermediates and how molecular chaperones enable productive folding. Real-time NMR spectroscopy, together with the development of advanced methods, is reviewed here and the potential these methods have to characterize intermediate states as well as interactions with molecular chaperone proteins at single-residue resolution is highlighted. Scope of review: Various chaperone interactions can guide the protein-folding reaction and thus are important for protein structure formation, stability, and activity of their substrates. Chaperone-assisted protein folding, characterization of intermediates, and their molecular interactions using real-time NMR spectroscopy will be discussed. Additionally, recent advances in NMR methods employed for characterization of high-energy intermediates will be discussed. Major conclusions: Real-time NMR combines high resolution with kinetic information of protein reactions, which can be employed not only for protein-folding studies and the characterization of folding intermediates but also to investigate the molecular mechanisms of assisted protein folding. General significance: Real-time NMR spectroscopy remains an effective tool to reveal structural details about the interaction between chaperones and transient intermediates. Methodologically, it provides in-depth understanding of how kinetic intermediates and their thermodynamics contribute to the protein-folding reaction. This review summarizes the most recent advances in this field. This article is part of a Special Issue titled Proline-directed Foldases: Cell Signaling Catalysts and Drug Targets. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Protein folding is the late step during the translation of genetic information into a biologically active polypeptide conformation. The biological macromolecules exist in different conformational states, where the native, well-structured state (N) is thermodynamically favored (Fig. 1). The folding of a protein in a given environment toward its final, thermodynamically favored conformation is governed by the primary amino acid sequence and only this conformation is biologically active. This traditional structure–function paradigm has been extended during the last 15 years by the following discoveries: (i) intrinsically disordered proteins can have a biological function [1]; (ii) the thermodynamically most stable protein structure can lack biological function [2]; and (iii) protein aggregates and well-structured amyloid fibrils are

☆ This article is part of a special issue titled Proline-directed Foldases: Cell Signaling Catalysts and Drug Targets. ⁎ Corresponding author at: Institut für Physik, Biophysik, Martin-Luther Universität Halle-Wittenberg, Betty-Heimann-Str. 7, Halle (Saale) D-06120, Germany. Tel.: +49 345 5528550; fax: +49 345 5527161. E-mail address: [email protected] (J. Balbach).

thermodynamically even more favored with respect to pathogenic function [3] (Fig. 1). Additionally, biological processes also depend on the passage through transition states or excited and high-energy states, which are often important, for example, during ligand binding, allosteric regulation, enzymatic catalysis, and protein folding [4,5]. Our current understanding of the energy landscapes of proteins was initiated by pioneering studies of the “new view of protein folding", and this has been achieved from both an experimental and a theoretical perspective [6–10]. The protein-folding process typically is quite fast for small, single-domain proteins [11–15], whereas this process is slower for larger or multidomain proteins, which follow a rugged funnel-shaped energy landscape to adopt the final functional state. During the slow-folding processes, long-life kinetic folding intermediates appear, which can lead to self-aggregation and malfunction of the protein in living cells [5,16, 17]. Therefore, nature has evolved specialized protein-folding “helpers", which prevent proteins from aggregating and/or speed up the folding process. Protein-folding intermediates expose hydrophobic patches recognized by the folding-helper enzymes to prevent unproductive aggregation [18–20]. There are three main classes of protein-folding accessory proteins found in nature. (i) Protein disulfide isomerases facilitate the shuffling

http://dx.doi.org/10.1016/j.bbagen.2014.12.003 0304-4165/© 2014 Elsevier B.V. All rights reserved.

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

2

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

Fig. 1. Energy landscape of different conformations of proteins. U represents the unfolded state, N the native state, and ‡ the transition state. Protein-folding catalysts such as PPIases reduce the kinetic barrier and favor the U → N reaction, whereas chaperones prevent formation of very stable aggregates. The expanded view of the profile on the right shows a more realistic view of the native state with additional intermediate high-energy states I1 and I2 that are accessible for the polypeptide chain due to thermal fluctuations.

of disulfide bonds until they achieve correct pairing. (ii) Peptidyl–prolyl cis–trans isomerases catalyze the slow interconversion of a Xaa-Pro peptide bond between the cis and trans conformation. Because of the partial double-bond character of the peptide bond, a high activation energy of about 80 kJ/mol has to be overcome resulting in folding time constants between 10 s and 100 s [21]. (iii) Molecular chaperones prevent protein aggregation by interacting with hydrophobic patches exposed by the folding substrate. By definition, molecular chaperones are a family of cellular proteins that assist non-covalent folding and unfolding, as well as assembly or disassembly of their substrates while they are not part of the final structure [22]. Thereby, they help in blocking alternative assembly or folding pathways that produce non-functional structures, e.g., by self-aggregation. Two principle chaperone classes can be defined: (i) foldases, which support the folding of proteins in an ATPdependent manner (e.g., GroEL/GroES or the DnaK/DnaJ/GrpE system), and (ii) holdases, which bind folding intermediates to prevent their aggregation (e.g., DnaJ or Hsp33) [23]. Protein disulfide isomerases and peptidyl–prolyl cis/trans-isomerases (PPIases) are well-studied examples of protein-folding catalysis [24,25]. In the energy landscape view of assisted protein folding, these classes of helper enzymes reduce the kinetic barrier toward the native state (dotted line in Fig. 1); thus, they speed up the productive reaction (green area in Fig. 1) with respect to the non-productive ones (red area in Fig. 1). The formation of disulfide bonds of proteins for example in the periplasm by molecular oxygen or in the eukaryotic endoplasmatic reticulum is far too slow to accompany protein folding on a second of minute timescale. Therefore, this process gets catalyzed by disulfide isomerases, which have been identified and studied in all three kingdoms of life. For PPIases, several catalytic mechanisms have been proposed [26], including (i) stabilization of a more apolar twisted prolyl bond by the hydrophobic enzyme environment, (ii) hydrogen bonding to the prolyl carbonyl oxygen by an enzyme bound water molecule, (iii) nucleophilic catalysis, (iv) protonation of the imide nitrogen, and (v) electrostatic transition-state stabilization. Proteins such as SurA [27], FkpA [28,29], trigger factor [30–32], and MtFKBP17 [33] comprise both PPIase and chaperone function located on two different domains. These two domains, however, act synergistically thereby leading to a protein-folding helper enzymes with high catalytic efficiency. SlyD, a member of the FK506 binding protein (FKBP) family, is also one of the well-characterized folding-helper enzymes present in Escherichia coli, harboring this dual function [34–37]. Protein-dependent biological processes are often accomplished via high-energy and/or short-lived intermediate states. Therefore, advanced experimental techniques are required to provide a comprehensive

insight into the conformations and dynamic behavior of these intermediate states at molecular resolution. Here we review recent progress in this direction for protein folding, assisted protein folding, and high-energy intermediates where NMR played a significant role in providing residue-specific information, structure, and dynamics of the respective proteins. NMR provides the dynamic behavior of molecules at high resolution over a wide range of timescales from picoseconds to days (Fig. 2). Therefore, NMR has emerged as a state-of-the-art technique to study the dynamics of proteins and protein reactions at molecular resolution. As discussed here, real-time NMR covers the timescale between milliseconds and hours. The principle experimental setup and analysis of 1D and 2D real-time NMR have been reviewed [38,39], as well as the different recent approaches to technically improve the time resolution [40], so these topics will not be covered in this review. Since the beginning of real-time NMR, the rapid sample-mixing devices have been improved to reduce dead-times below 100 ms [41] or even tens of milliseconds [42], and the fastest protein-folding events directly observed by NMR after rapid mixing occurred on a 100 ms timescale [43]. For molecular resolution, two- and multidimensional NMR experiments are required, and currently protein reactions not faster than 10−2 s−1 can be characterized [40]. At equilibrium and under suitable conditions (temperature, denaturant concentrations, and pH value), the protein chain fluctuates between the native-state, high-energy folding intermediates, and the unfolded state. These fluctuations down to a microsecond timescale can be followed by NMR relaxation and relaxation dispersion experiments [44–46]. The first high-resolution structures of short-lived onpathway intermediates at these timescales could be determined for the Fyn SH3 domain [47]. Amide proton exchange adds a further level of information, because this process provides insights not only regarding the global thermodynamic stability of the protein but also regarding the local stability of each individual residue [48]. In combination with real-time NMR, this method allows the assessment of the formation of single hydrogen bonds during late steps in protein folding and sheds light on how networks of hydrogen bonds thermodynamically couple the stability of two-domain proteins [49]. In this review article, we will discuss the recent developments in real-time NMR of proteins and present specific examples for the characterization of short-lived protein-folding intermediates and chaperoneassisted protein folding.

2. Advances in multidimensional real-time NMR spectroscopy Progress in NMR spectroscopy in terms of method development and innovative technologies during the last 10 years permits the analysis of large and complex biomolecules, including unstructured proteins and enables researchers to follow time-resolved protein reactions at high resolution. These developments and their principle requirements will be briefly discussed in this section. Bacterial systems are the method of choice for labeling pro- and eukaryotic proteins with NMR-active isotopes, typically 2H/13C/15 N. If post-translational modifications or a cellular folding assistance are required in eukaryotic cells, yeast expression systems (Pichia pastoris or Kluyveromyces lactis [50]) or baculovirus-mediated expression systems in mammalian cells [51,52] are most popular. For the latter, insect cells are often used, especially to allow the correct post-translational modifications to occur [53,54] and CHO or HEK 293 cells can be used for the same purpose [55]. Cell-free protein production is the method of choice for high-throughput protein production and selective incorporation of isotope-labeled amino acids into a target protein for both solution and solid-state NMR [56]. Combining multidimensional NMR, novel pulse sequences and the development of various labeling techniques has allowed high-resolution NMR analyses of supramolecular systems, e.g., a SecA ATPase/peptide complex (204 kDa) [57], the 20S proteasome

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

3

Fig. 2. NMR timescale and the respective method/NMR parameter sensitivity in the study of dynamic motion and reactions of biomacromolecules. Red represents topics covered by this review.

[58,59], or mega-dalton protein complexes and beyond [60], to be realized. Another advantage of solution NMR spectroscopy compared to other high-resolution structural biology methods is the possibility to study time-resolved protein reactions. To structurally characterize transient intermediate states, e.g., those appearing during protein folding, fast multidimensional techniques are required. A variety of recent developments have succeeded in this direction by speeding up normal timedomain sampling and the pulsing repetition rate (see reviews [61, 62]). Optimized spectral aliasing [63], spectral reconstruction algorithms that can handle sparse, non-uniformly sampled data sets [64–66], Hadamard-encoding techniques [67], and spatially encoded, single-scan methods [68,69] use sparse data sampling for fast recording of real-time NMR data. The advent of non-uniform sampling paved the way into a new era of multidimensional NMR spectroscopy in short time (more details can be found in reviews [70–72]). On the other hand, fast-pulsing techniques such as flip-back approaches [73,74], BEST [63,75–78], and SOFAST [74,79] sequences use significantly shorter inter-scan delays. For this purpose, longitudinal 1H relaxation is enhanced in case of BEST and SOFAST by manipulating the protons by shaped and band-selective radio frequency pulses. These pulses will only excite the protons of interest (e.g., backbone amides), and all other protons remain non-perturbed. This causes strongly reduced longitudinal 1H relaxation times of the observed nuclei because of dipolar interactions between both sets of protons. SOFAST-HMQC experiments additionally utilize the Ernst-angle excitation to increase the 1H steady-state polarization by reducing both the effective flip angle of the 1H excitation and the preceding longitudinal relaxation delay for an optimal signal-to-noise ratio per experimental time unit. SOFASTHMQC has emerged as the best 2D real-time NMR technique to provide the maximum sensitivity in the shortest time-span. Using this technique, complete 2D spectra of proteins or DNA can be achieved in less than 5 s (Fig. 3), which is currently the time-limit for in-depth analysis of structure formation. Fast pulsing can be combined with various other approaches, e.g., with sparse sampling employing spatially encoded single-scan acquisition, termed ultraSOFAST [80,81]. Conventional phase cycling in order to remove experimental artifacts is typically applied between successive 2D data acquisitions in real-time 2D SOFAST NMR [82]. In the BEST-TROSY experiment, otherwise lost 1H polarization gets converted into 15N polarization in the subsequent scan to significantly increase the signal-to-noise ratio [76]. The conformational dynamics along the protein chain is also accessible by fast-pulsing methods. HET-SOFAST or HET-BEST samples the local compactness and fast timescale motions by selectively saturating the solvent or non-detected protons before the SOFAST or BEST sequence [78]. Conformational dynamics on the μs to ms timescale elucidated by transversal relaxation includes R2-BEST-HSQC or R2-BEST-TROSY [63,75,76,83, 84], and for the characterization of the overall size and oligomerization state of protein states, BEST-TRACT [85] is useful.

Depending upon the experimental setup, fast-pulsing conditions allow 2- to 10-fold sensitivity to be achieved using SOFAST and BEST. Utilizing the same methods, 3D data collection can be reduced from days to less than 1 h. For more details about the method, readers may refer to the references mentioned above and a recent review by Rennella and Brutscher [40].

Fig. 3. Two-dimensional real-time NMR analysis of ubiquitin and characterization of a protein-folding intermediate. (A) SOFAST real-time 2D NMR timescale. (B) SOFAST real-time 2D NMR spectra recorded within the second to minute time frame. The enlarged spectrum was recorded in an experimental time frame of 4 second with 0.2 mM 15N-ubiquitin at 18.8 T magnetic field strength and a cryogenically cooled probe. The figure was taken with permission from [82] copyright (2007) National Academy of Sciences, USA.

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

4

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

3. Folding intermediates and assisted protein folding studied by real-time NMR Traditionally, protein-folding intermediates are identified when analyzing folding kinetics using optical spectroscopy methods. These techniques give a wealth of information about the various timescales of interconversion of the unfolded, intermediate, and native states and their thermodynamic energy landscapes. Although a time resolution down to nanoseconds [86,87] can be achieved, these studies are limited in their structural resolution. The role of individual residues can be revealed by point mutations, but a comprehensive study is very time consuming with many obstacles to overcome. On the other hand, NMR spectroscopy gives molecular resolution of proteins in solution but is an intrinsically slow method. As discussed in the previous section, current time-resolved protein NMR at high resolution still requires at best several seconds of data acquisition. Real-time NMR spectroscopy has been applied to several biological systems including, for example, structural studies of transient proteinfolding intermediates (see above), histone deacetylase and acetyltransferase activities [88], RNA folding [89], intermediates and labile products of enzymes [90], and multi-phosphorylation of intrinsically disordered protein domains [91]. Real-time NMR application to protein folding has been extensively reviewed [38,89,92–95]. Due to the scope of this special journal issue, a specific example of assisted protein folding monitored by real-time NMR spectroscopy will be discussed (see below). With the recent advances in NMR spectroscopy it is now possible to characterize short-lived intermediate states, e.g., by assigning their backbone amides. Brutscher and co-workers achieved this for β2microglobulin by analyzing 3D-BEST-TROSY HNCA and HNCO spectra of an intermediate state (I). During the protein refolding process this state converts into the native state (N). This protein reaction occurs during data acquisition of a single 3D data set and therefore the resonances of both the I and the N states are included in the real-time 3D spectra. Acquisition time should match the refolding kinetics to enhance the signal-noise ratio and to differentiate the cross peaks of I and N according to their line shapes in the indirect dimension [83,96]. To obtain the pure I state spectrum, the N state resonances were subtracted from the real-time 3D data by recording another 3D data set directly after refolding has finished. The latter data set contains only resonances of N. Before subtraction, an apodization function representing the folding kinetics of N needs to be applied for this purpose [83,95,97]. The thus achieved assignments of the β2-microglobulin folding intermediate could be used to identify the already present secondary structures as well as the local dynamics in comparison to the native state [83]. A similar approach was applied to study in considerable detail a protein-folding intermediate populated during the refolding of ribonuclease T1 (RNase T1) from Aspergillus oryzae, which is a single-domain 104 amino acid protein with four proline residues. Proline residues 39 and 55 are in the cis conformation in the native state and are the main retarding determinants of the folding mechanism [98]. Introduction of two mutations S54G/P55N led to a simple overall folding mechanism and showed only one long-lived native-like intermediate (I39t) having proline 39 in the not-yet-native trans conformation. In vitro refolding of denatured S54G/P55N RNase T1 in highly concentrated GdmCl solutions follows the folding pathway U → I39t → N39c. The folding intermediate I39t could be structurally characterized by real-time NMR experiments with molecular resolution [77,97,99]. The lifetime of this transient state is about 5 h as shown by fluorescence and 1D 1H NMR spectroscopy. Therefore, a conventional HNCA experiment could not be used but a BEST-HNCA for 5 h with a reduced inter-scan delay time from 1 s to 200 ms was sufficient for NMR resonance assignments. This 3D spectrum contains resonances of both I39t and the N state. The resonances of the N state can be assigned by recoding conventional 3D HNCA, HNCACB, HN(CO)CACB, 15N edited NOESY and TOCSY experiments after the refolding process has finished. These assignments

could be used in the kinetic 3D BEST-HNCA to identify the resonances of the N state. All additional cross peaks belong to I39t. Furthermore, cross peaks of I39t differ as mentioned above in their line shape [96,97] because the population of I39t decreases and the population of N increases during recording of the kinetic 3D BEST-HNCA (Fig. 4A). This change in population because of the ongoing protein-folding reaction during data acquisition can be observed, e.g., for the amide backbone resonance of A19 in Fig. 4C. Strips of the “sequential walk” from A19 to K25 is given in Fig. 4B for I39t and in Fig. 4D for the N state as representative examples from the kinetic 3D BEST-HNCA. From the derived NMR resonance assignments of I39t, its secondary structure elements can be determined by the backbone chemical shift index (Fig. 4E) and color-coded on the protein structure (Fig. 4F) [77]. The majority of residues in I39t shows the chemical shift index of the native state and therefore has formed already the native conformation before the rate-limiting isomerization of Pro39 from the trans to the cis conformation occurs. Residues in close proximity to Pro39 as well as the C-terminal half of the only α-helix of S54G/P55N RNase T1 are in a not-yet-native conformation. An earlier study showed by H/D exchange and NOE measurements that the α-helix has already formed [97] but differs from the native-like structure most probably by having a different orientation relative to Pro39. The hydrogen–deuterium (H/D) exchange is a powerful method to measure the local thermodynamic backbone stability because the rate of exchange of amide protons drop by orders of magnitude in ordered secondary structures resulting in high protection factors against exchange with solvent protons [48]. For the five-repeat protein p19INK4d, we found a good correlation between these protection factors in the N state and the structured regions in an on-pathway intermediate [100]. The N-terminal ankyrin repeats 1 and 2 are unstructured in the intermediate and less protected against by H/D exchange in the N state, whereas repeats 3–5 are structured in the intermediate and are highly protected. Therefore, repeats 3–5 form first as a scaffold for the subsequent assembly of repeats 1 and 2. H/D exchange can also be used to directly determine already formed hydrogen bonds in protein-folding intermediates. This was successful for example in the case of the gene-3-protein from the tip of filamentous phage controlling their infectivity via the TolA receptor [49]. Trans to cis isomerization of Pro 312 controls the assembly of the N1 and N2 domain of the gene-3-protein representing the “timer” for phage infection. Real-time NMR revealed the residues that change their conformation during the rate-limiting assembly step. They were mainly located in the N1 domain of the gene-3-protein and at the interface between the two domains. A competition between H/D exchange and this final folding step disclosed which hydrogen bonds form last in the hinge region between the two domains and thus mediate domain assembly. The latter directly controls phage infectivity. To increase the time resolution for H/D exchange of proteins, the HETex-SOFAST NMR method is suitable [79]. Recently, it has been extended to BEST readout sequences without losing sensitivity and with better resolution, while allowing extraction of quantitative exchange rates from HETex NMR measurements. The method was successfully applied to α-synuclein, human ubiquitin, E. coli heat-shock protein CspA, and amyloidogenic protein β2-microglobulin [101]. Under native conditions, in addition to the native, fully folded protein conformation, high-energy states can exist, which are still accessible for the peptide chain by thermal fluctuations at equilibrium (closeup in Fig. 1). As discussed above, these states can be analyzed by NMR relaxation methods. Another achievement in NMR spectroscopy has been the characterization of protein-folding intermediates under high pressure. According to the volume of the respective protein state in its given environment, a pressure change can shift the populations at equilibrium [102]. This might have a distinct advantage over the other denaturation methods in terms of reversibility. Rare “excited” conformational states, for example, intermediates on the pathway to amyloid fibrils that have a relative population of as small as 0.0005 at ambient

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

(A)

5

(B)

N

time [h]

(C)

(D) [ time

s] 100

(F)

(E) chemical shift shift index chemical index(C(C) )

8 6

I39t

4 2 0 -2 -4

N

-6 -8 0

20

40

60

80

100

residue number Fig. 4. Real-time NMR characterization of the protein-folding intermediate I39t of S54G/P55N RNase T1. (A) Time course of the population of the native state (N) and I39t together with the recorded NMR experiments. (B) and (D): backbone resonance assignment by “sequential walking” through the 3D BEST-HNCA stripes of residues A19-K25. (C) NMR intensity changes reporting the protein-folding reaction in sequential 2D 1H-15N TROSY HSQC spectra. (E) The chemical shift index reveals the secondary structural elements (below −2 ppm for β-strands and above 2 ppm for α-helices) and the black and gray bars represent the corresponding elements in the final native state. (F) Color-coding of the backbone chemical shifts of I39t on the structure of RNase T1. Blue: native chemical shifts and gold: not-yet-native chemical shifts. The figure was reprinted with permission from [77] copy right American Chemical Society (2011).

pressure, can be stabilized by pressure and thus detected [103,104]. Other advantages of high-pressure NMR for the study of protein folding, protein aggregation, and ligand interactions at molecular resolution have been reviewed [105]. Recently, Heuert et al. have developed an NMR probe for pressurejump experiments up to 250 bars and 3 ms jump time [106]. This probe head is able to perform under a computer-controlled pressure step from high pressure of ~ 10 MPa to ambient pressure followed by 1D 1H NMR spectroscopy. However, for protein NMR, a much higher pressure step and involvement of advanced NMR methods are required. The Kalbitzer group has developed a microprocessor-controlled pressure-jump unit to analyze protein folding under pressure. Using this methodology, they were able to introduce fast and strong pressure changes at any point in the NMR pulse sequences where, in a sample tube, repetitive pressure change of 80 MPa can be introduce within

less than 30 ms [107]. Two kinds of experiments are possible: (i) pressure perturbation transient state spectroscopy (PPTSS), which allows the determination of the rates, activation energies, and activation volumes between different protein states, e.g., during protein folding or ligand binding, and (ii) pressure perturbation state correlation spectroscopy (PPSCS), which allows the transfer of the measured properties of a pressure-stabilized state to another protein state for a proper readout. The slow refolding of S54G/P55N RNase T1 retarded by the trans to cis peptidyl–prolyl isomerization of the Y38-P39 peptide bond, as well as the characterization of the intermediate state I39t using real-time NMR, have been discussed in the previous section. PPIases are ubiquitous catalysts in the cell, which speed up such slow refolding steps [108]. Sensitive-to-lysis protein D (SlyD), an FKBP family member from the bacterial cytoplasm contains a PPIase domain and an inserted “in-flap” (IF domain) chaperone domain between β-strand 2 and 3

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

6

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

SlyD (Enzym)

chaperone

RNase T1 (Substrat)

PPIase

E+S

k1 k-1

ES

k2

E+P

Fig. 5. Real-time NMR chemical shift mapping of a transient enzyme-substrate complex. Catalysis of the refolding of S54G/P55N RNase T1 by SlyD follows a Michaelis–Menten mechanism. Residues involved in the protein-protein interaction in the enzyme-substrate (ES) complex derived from real-time NMR experiments are color coded in red on the structures of each protein (protein database codes 2K8I and 9RNT). The figure was reprinted with permission from [77], copyright American Chemical Society (2011).

(Fig. 5). The molecular mechanism related to the distinct interplay of the two domains to make SlyD as an efficient PPIase with very high catalytic efficiency has been recently reviewed [109,110]. The S54G/P55N RNase T1 is a substrate of SlyD and its catalyzed refolding can be described by a Michaelis–Menten mechanism [35]. We could structurally characterize the transient enzyme-substrate recognition by real-time NMR spectroscopy (Fig. 5) [77]. The backbone NMR chemical shifts of transient I39t in the uncatalyzed refolding reaction could be assigned (Fig. 4). During catalysis, SlyD binds to I39t and forms the enzymesubstrate complex ([ES] in Fig. 5). Therefore, the experiments shown in Fig. 4 were repeated but in the presence of SlyD and residues interacting with SlyD are colored red in Fig. 5. This causes changes in the chemical shifts of I39t of those residues, which interact with SlyD. The majority of these residues correspond to residues that are in a not-yet-native conformation in I39t because of the trans conformation of Pro39. The same analysis was possible for SlyD, where backbone assignments and the NMR structure are known [37]. Nearly all residues of the chaperone domain of SlyD change chemical shift positions as well as residues of the active site of the PPIase domain (Fig. 5). In addition to these structural features of the ES complex, the prerequisites of the Michaelis–Menten mechanism could be confirmed. It implies that the rate constants k1 and k-1 are much higher compared to k2. Indeed, the real-time NMR experiments revealed that the changes of the NMR resonances between E and ES occurred in the fast exchange regime on the NMR chemical shift timescale. The transition of ES to P, characterized by k2, occurs on in the slow exchange regime on the NMR chemical shift timescale because separate chemical shifts could be observed for ES and P. Finally, SlyD and fully folded S54G/P55N RNase T1 showed a weak interaction, which could be monitored by NMR titration of SlyD with the folded substrate, corresponding to an enzyme-product complex. Because of this tight interaction between SlyD and its substrate, we speculate that from the various possibilities to catalyze a peptidyl– prolyl bond isomerization [26], exclusion of water accessibility to Pro 39 in RNase T1 (S54G/P55N) and a hydrophobic environment provided by SlyD represents the most likely mechanism [77]. In the energy landscape view (Fig. 1), SlyD reduces the kinetic barrier of refolding of S54G/ P55N RNase T1 by this mechanism. Various biochemical studies highlighted the synergistic interplay of the chaperone and PPIase domains of SlyD [35,36,109–111]. The first step during protein-folding catalysis of SlyD is the unspecific binding of the chaperone domain to the partly folded substrate, which even

overrides the specificity of the catalytic center in the PPIase domain. As a classical chaperone, SlyD also prevents protein aggregation (Fig. 1) by binding with its chaperone domain to hydrophobic and aggregation-prone segments [35,36]. Real-time NMR revealed that almost exclusively residues of the chaperone domain interact with the aggregation-prone substrate and that binding occurs already during the lag phase of, e.g., insulin aggregation, which almost completely suppresses aggregation [37]. During the second step of catalysis, the bound substrate gets close to the catalytic center of SlyD located in the PPIase domain by a continuous opening and closing of the chaperone and PPIase domain. This allows the enzyme to sample various orientations of the folding substrate relative to the catalytic center before isomerization of the prolyl peptide bond and release of the substrate occurs [109,110]. That long-lived protein-folding intermediates, the classical substrates for chaperones, are prone to oligomerization has also been quantified recently by real-time NMR in combination with other biophysical methods for beta-2-micorglobulin [40]. 4. Concluding remarks The rapid progress in the method development of real-time NMR together with advanced NMR methods for fast data acquisition and isotope labeling schemes form the basis that allow protein reactions to be studied in real-time and in molecular detail. This deepens our understanding of protein-folding, enzyme catalysis, and high-energy protein states, which is difficult to achieve with other biophysical or structural biological methods. Acknowledgments This research was supported by grants from the DFG (GRK 1026, SFB TRR102), the BMBF (ProNet-T3), and ERDF by the EU for significant investigations into the NMR facility. We thank Gary Sawers for very helpful discussions. References [1] H.J. Dyson, P.E. Wright, Intrinsically unstructured proteins and their functions, Nat. Rev. Mol. Cell Biol. 6 (2005) 197–208. [2] B. Eckert, A. Martin, J. Balbach, F.X. Schmid, Prolyl isomerization as a molecular timer in phage infection, Nat. Struct. Mol. Biol. 12 (2005) 619–623.

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx [3] T. Eichner, S.E. Radford, A diversity of assembly mechanisms of a generic amyloid fold, Mol. Cell 43 (2011) 8–18. [4] A.J. Baldwin, L.E. Kay, NMR spectroscopy brings invisible protein states into focus, Nat. Chem. Biol. 5 (2009) 808–814. [5] C.M. Dobson, Principles of protein folding, misfolding and aggregation, Semin. Cell Dev. Biol. 15 (2004) 3–16. [6] C.M. Dobson, A. Sali, M. Karplus, Protein folding: a perspective from theory and experiment, Angew. Chem. Int. Ed. Engl. 37 (1998) 868–893. [7] K.A. Dill, J.L. MacCallum, The protein-folding problem, 50 years on, Science 338 (2012) 1042–1046. [8] D.U. Ferreiro, E.A. Komives, P.G. Wolynes, Frustration in biomolecules, Q. Rev. Biophys. (2014) 1–79. [9] R.L. Baldwin, G.D. Rose, Is protein folding hierarchic? II. Folding intermediates and transition states, Trends Biochem. Sci. 24 (1999) 77–83. [10] R.L. Baldwin, G.D. Rose, Is protein folding hierarchic? I. Local structure and peptide folding, Trends Biochem. Sci. 24 (1999) 26–33. [11] W.A. Eaton, V. Munoz, S.J. Hagen, G.S. Jas, L.J. Lapidus, E.R. Henry, J. Hofrichter, Fast kinetics and mechanisms in protein folding, Annu. Rev. Biophys. Biomol. Struct. 29 (2000) 327–359. [12] N. Ferguson, A.R. Fersht, Early events in protein folding, Curr. Opin. Struct. Biol. 13 (2003) 75–81. [13] A.R. Fersht, Transition-state structure as a unifying basis in protein-folding mechanisms: contact order, chain topology, stability, and the extended nucleus mechanism, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 1525–1529. [14] S.E. Jackson, How do small single-domain proteins fold? Fold. Des. 3 (1998) R81–R91. [15] T. Schindler, M. Herrler, M.A. Marahiel, F.X. Schmid, Extremely rapid protein folding in the absence of intermediates, Nat. Struct. Biol. 2 (1995) 663–673. [16] C.M. Dobson, Protein folding and misfolding, Nature 426 (2003) 884–890. [17] T.R. Jahn, S.E. Radford, The yin and yang of protein folding, FEBS J. 272 (2005) 5962–5970. [18] B. Bukau, E. Deuerling, C. Pfund, E.A. Craig, Getting newly synthesized proteins into shape, Cell 101 (2000) 119–122. [19] S. Walter, J. Buchner, Molecular chaperones–cellular machines for protein folding, Angew. Chem. Int. Ed. Engl. 41 (2002) 1098–1113. [20] J.C. Young, V.R. Agashe, K. Siegers, F.U. Hartl, Pathways of chaperone-mediated protein folding in the cytosol, Nat. Rev. Mol. Cell Biol. 5 (2004) 781–791. [21] J. Balbach, F.X. Schmid, Proline isomerization and its catalysis in protein folding, in: R.H. Pain (Ed.), Mechanisms of Protein Folding, University Press, Oxford, 2000, pp. 212–237. [22] J. Buchner, S. Walter, Analysis of chaperone function in vitro, in: J. Buchner, T. Kiefhaber (Eds.),Handbook of Protein Folding, vol. 1, Wiley-VCH, Weinheim, 2004, pp. 162–196. [23] R.J. Ellis, Molecular chaperones: assisting assembly in addition to folding, Trends Biochem. Sci. 31 (2006) 395–401. [24] J.F. Collet, J. Riemer, M.W. Bader, J.C. Bardwell, Reconstitution of a disulfide isomerization system, J. Biol. Chem. 277 (2002) 26886–26892. [25] G. Fischer, B. Wittmann-Liebold, K. Lang, T. Kiefhaber, F.X. Schmid, Cyclophilin and peptidyl–prolyl cis–trans isomerase are probably identical proteins, Nature 337 (1989) 476–478. [26] J. Fanghänel, G. Fischer, Insights into the catalytic mechanism of peptidyl prolyl cis/ trans isomerases, Front. Biosci. 9 (2004) 3453–3478. [27] P.E. Rouviere, C.A. Gross, SurA, a periplasmic protein with peptidyl–prolyl isomerase activity, participates in the assembly of outer membrane porins, Genes Dev. 10 (1996) 3170–3182. [28] K. Ramm, A. Pluckthun, High enzymatic activity and chaperone function are mechanistically related features of the dimeric E. coli peptidyl–prolyl–isomerase FkpA, J. Mol. Biol. 310 (2001) 485–498. [29] F.A. Saul, J.P. Arie, B. Vulliez-le Normand, R. Kahn, J.M. Betton, G.A. Bentley, Structural and functional studies of FkpA from Escherichia coli, a cis/trans peptidyl– prolyl isomerase with chaperone activity, J. Mol. Biol. 335 (2004) 595–608. [30] T. Hesterkamp, B. Bukau, The Escherichia coli trigger factor, FEBS Lett. 389 (1996) 32–34. [31] C. Scholz, G. Stoller, T. Zarnt, G. Fischer, F.X. Schmid, Cooperation of enzymatic and chaperone functions of trigger factor in the catalysis of protein folding, EMBO J. 16 (1997) 54–58. [32] G. Stoller, K.P. Rucknagel, K.H. Nierhaus, F.X. Schmid, G. Fischer, J.U. Rahfeld, A ribosome-associated peptidyl–prolyl cis/trans isomerase identified as the trigger factor, EMBO J. 14 (1995) 4939–4948. [33] R. Suzuki, K. Nagata, F. Yumoto, M. Kawakami, N. Nemoto, M. Furutani, K. Adachi, T. Maruyama, M. Tanokura, Three-dimensional solution structure of an archaeal FKBP with a dual function of peptidyl prolyl cis–trans isomerase and chaperone-like activities, J. Mol. Biol. 328 (2003) 1149–1160. [34] L. Martino, Y. He, K.L. Hands-Taylor, E.R. Valentine, G. Kelly, C. Giancola, M.R. Conte, The interaction of the Escherichia coli protein SlyD with nickel ions illuminates the mechanism of regulation of its peptidyl–prolyl isomerase activity, FEBS J. 276 (2009) 4529–4544. [35] C. Scholz, B. Eckert, F. Hagn, P. Schaarschmidt, J. Balbach, F.X. Schmid, SlyD proteins from different species exhibit high prolyl isomerase and chaperone activities, Biochemistry 45 (2006) 20–33. [36] C. Scholz, P. Schaarschmidt, A.M. Engel, H. Andres, U. Schmitt, E. Faatz, J. Balbach, F.X. Schmid, Functional solubilization of aggregation-prone HIV envelope proteins by covalent fusion with chaperone modules, J. Mol. Biol. 345 (2005) 1229–1241. [37] U. Weininger, C. Haupt, K. Schweimer, W. Graubner, M. Kovermann, T. Bruser, C. Scholz, P. Schaarschmidt, G. Zoldak, F.X. Schmid, J. Balbach, NMR solution structure

[38] [39]

[40]

[41]

[42]

[43]

[44]

[45] [46]

[47]

[48] [49]

[50]

[51] [52]

[53] [54] [55] [56]

[57]

[58] [59] [60]

[61] [62] [63]

[64]

[65]

[66] [67]

[68] [69] [70]

7

of SlyD from Escherichia coli: spatial separation of prolyl isomerase and chaperone function, J. Mol. Biol. 387 (2009) 295–305. M. Zeeb, J. Balbach, Protein folding studied by real-time NMR spectroscopy, Methods 34 (2004) 65–74. M. Zeeb, J. Balbach, Kinetic protein folding studies using NMR spectroscopy, in: J. Buchner, T. Kiefhaber (Eds.),Handbook of protein folding, vol. 1, Wiley-VCH, Weinheim, 2004, pp. 536–572. E. Rennella, T. Cutuil, P. Schanda, I. Ayala, F. Gabel, V. Forge, A. Corazza, G. Esposito, B. Brutscher, Oligomeric states along the folding pathways of beta2-microglobulin: kinetics, thermodynamics, and structure, J. Mol. Biol. 425 (2013) 2722–2736. J.G. Bann, J. Pinkner, S.J. Hultgren, C. Frieden, Real-time and equilibrium (19)F-NMR studies reveal the role of domain-domain interactions in the folding of the chaperone PapD, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 709–714. K.H. Mok, T. Nagashima, I.J. Day, J.A. Jones, C.J. Jones, C.M. Dobson, P.J. Hore, Rapid sample-mixing technique for transient NMR and photo-CIDNP spectroscopy: applications to real-time protein folding, J. Am. Chem. Soc. 125 (2003) 12484–12492. K.H. Mok, L.T. Kuhn, M. Goez, I.J. Day, J.C. Lin, N.H. Andersen, P.J. Hore, A preexisting hydrophobic collapse in the unfolded state of an ultrafast folding protein, Nature 447 (2007) 106–109. A.G. Palmer 3rd, C.D. Kroenke, J.P. Loria, Nuclear magnetic resonance methods for quantifying microsecond-to-millisecond motions in biological macromolecules, Methods Enzymol. 339 (2001) 204–238. M. Zeeb, M.H. Jacob, T. Schindler, J. Balbach, 15N relaxation study of the cold shock protein CspB at various solvent viscosities, J. Biomol. NMR 27 (2003) 221–234. M. Tollinger, N.R. Skrynnikov, F.A. Mulder, J.D. Forman-Kay, L.E. Kay, Slow dynamics in folded and unfolded states of an SH3 domain, J. Am. Chem. Soc. 123 (2001) 11341–11352. P. Neudecker, P. Robustelli, A. Cavalli, P. Walsh, P. Lundstrom, A. Zarrine-Afsar, S. Sharpe, M. Vendruscolo, L.E. Kay, Structure of an intermediate state in protein folding and aggregation, Science 336 (2012) 362–366. Y. Bai, T.R. Sosnick, L. Mayne, S.W. Englander, Protein folding intermediates: native-state hydrogen exchange, Science 269 (1995) 192–197. U. Weininger, R.P. Jakob, B. Eckert, K. Schweimer, F.X. Schmid, J. Balbach, A remote prolyl isomerization controls domain assembly via a hydrogen bonding network, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 12335–12340. T. Sugiki, O. Ichikawa, M. Miyazawa-Onami, I. Shimada, H. Takahashi, Isotopic labeling of heterologous proteins in the yeast Pichia pastoris and Kluyveromyces lactis, Methods Mol. Biol. 831 (2012) 19–36. Y.C. Hu, Baculoviral vectors for gene delivery: a review, Curr. Gene Ther. 8 (2008) 54–65. T.A. Kost, J.P. Condreay, R.S. Ames, S. Rees, M.A. Romanos, Implementation of BacMam virus gene delivery technology in a drug discovery setting, Drug Discov. Today 12 (2007) 396–403. T.A. Kost, J.P. Condreay, D.L. Jarvis, Baculovirus as versatile vectors for protein expression in insect and mammalian cells, Nat. Biotechnol. 23 (2005) 567–575. K. Saxena, A. Dutta, J. Klein-Seetharaman, H. Schwalbe, Isotope labeling in insect cells, Methods Mol. Biol. 831 (2012) 37–54. Y. Durocher, M. Butler, Expression systems for therapeutic glycoprotein production, Curr. Opin. Biotechnol. 20 (2009) 700–707. S. Sobhanifar, S. Reckel, F. Junge, D. Schwarz, L. Kai, M. Karbyshev, F. Lohr, F. Bernhard, V. Dötsch, Cell-free expression and stable isotope labelling strategies for membrane proteins, J. Biomol. NMR 46 (2010) 33–43. I. Gelis, A.M. Bonvin, D. Keramisanou, M. Koukaki, G. Gouridis, S. Karamanou, A. Economou, C.G. Kalodimos, Structural basis for signal-sequence recognition by the translocase motor SecA as determined by NMR, Cell 131 (2007) 756–769. R. Sprangers, L.E. Kay, Quantitative dynamics and binding studies of the 20S proteasome by NMR, Nature 445 (2007) 618–622. M.P. Latham, A. Sekhar, L.E. Kay, Understanding the mechanism of proteasome 20S core particle gating, Proc. Natl. Acad. Sci. U. S. A. 111 (2014) 5532–5537. A. Mainz, T.L. Religa, R. Sprangers, R. Linser, L.E. Kay, B. Reif, NMR spectroscopy of soluble protein complexes at one mega-dalton and beyond, Angew. Chem. Int. Ed. Engl. 52 (2013) 8746–8751. I.C. Felli, B. Brutscher, Recent advances in solution NMR: fast methods and heteronuclear direct detection, ChemPhysChem 10 (2009) 1356–1368. R. Freeman, E. Kupce, New methods for fast multidimensional NMR, J. Biomol. NMR 27 (2003) 101–113. E. Lescop, P. Schanda, B. Brutscher, A set of BEST triple-resonance experiments for time-optimized protein resonance assignment, J. Magn. Reson. 187 (2007) 163–169. B.E. Coggins, R.A. Venters, P. Zhou, Radial sampling for fast NMR: concepts and practices over three decades, Prog. Nucl. Magn. Reson. Spectrosc. 57 (2010) 381–419. K. Kazimierczuk, J. Stanek, A. Zawadzka-Kazimierczuk, W. Kozminski, Random sampling in multidimensional NMR spectroscopy, Prog. Nucl. Magn. Reson. Spectrosc. 57 (2010) 420–434. M. Mobli, J.C. Hoch, Maximum entropy spectral reconstruction of non-uniformly sampled data, Concepts Magn. Reson. Part A Bridg. Educ. Res. 32A (2008) 436–448. E. Lescop, R. Rasia, B. Brutscher, Hadamard amino-acid-type edited NMR experiment for fast protein resonance assignment, J. Am. Chem. Soc. 130 (2008) 5014–5015. M. Mishkovsky, L. Frydman, Principles and progress in ultrafast multidimensional nuclear magnetic resonance, Annu. Rev. Phys. Chem. 60 (2009) 429–448. A. Tal, L. Frydman, Single-scan multidimensional magnetic resonance, Prog. Nucl. Magn. Reson. Spectrosc. 57 (2010) 241–292. S.G. Hyberts, H. Arthanari, S.A. Robson, G. Wagner, Perspectives in magnetic resonance: NMR in the post-FFT era, J. Magn. Reson. 241 (2014) 60–73.

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003

8

A. Kumar, J. Balbach / Biochimica et Biophysica Acta xxx (2014) xxx–xxx

[71] S.G. Hyberts, H. Arthanari, G. Wagner, Applications of non-uniform sampling and processing, Top. Curr. Chem. 316 (2012) 125–148. [72] M. Mobli, M.W. Maciejewski, A.D. Schuyler, A.S. Stern, J.C. Hoch, Sparse sampling methods in multidimensional NMR, Phys. Chem. Chem. Phys. 14 (2012) 10835–108343. [73] H.S. Atreya, T. Szyperski, G-matrix Fourier transform NMR spectroscopy for complete protein resonance assignment, Proc. Natl. Acad. Sci. U. S. A. 101 (2004) 9642–9647. [74] T. Diercks, M. Daniels, R. Kaptein, Extended flip-back schemes for sensitivity enhancement in multidimensional HSQC-type out-and-back experiments, J. Biomol. NMR 33 (2005) 243–259. [75] J. Farjon, J. Boisbouvier, P. Schanda, A. Pardi, J.P. Simorre, B. Brutscher, Longitudinalrelaxation-enhanced NMR experiments for the study of nucleic acids in solution, J. Am. Chem. Soc. 131 (2009) 8571–8577. [76] A. Favier, B. Brutscher, Recovering lost magnetization: polarization enhancement in biomolecular NMR, J. Biomol. NMR 49 (2011) 9–15. [77] C. Haupt, R. Patzschke, U. Weininger, S. Groger, M. Kovermann, J. Balbach, Transient enzyme-substrate recognition monitored by real-time NMR, J. Am. Chem. Soc. 133 (2011) 11154–11162. [78] P. Schanda, V. Forge, B. Brutscher, HET-SOFAST NMR for fast detection of structural compactness and heterogeneity along polypeptide chains, Magn. Reson. Chem. 44 (2006) S177–S184 (Spec No). [79] P. Schanda, E. Kupce, B. Brutscher, SOFAST-HMQC experiments for recording twodimensional heteronuclear correlation spectra of proteins within a few seconds, J. Biomol. NMR 33 (2005) 199–211. [80] M. Gal, T. Kern, P. Schanda, L. Frydman, B. Brutscher, An improved ultrafast 2D NMR experiment: towards atom-resolved real-time studies of protein kinetics at multi-Hz rates, J. Biomol. NMR 43 (2009) 1–10. [81] M. Gal, P. Schanda, B. Brutscher, L. Frydman, UltraSOFAST HMQC NMR and the repetitive acquisition of 2D protein spectra at Hz rates, J. Am. Chem. Soc. 129 (2007) 1372–1377. [82] P. Schanda, V. Forge, B. Brutscher, Protein folding and unfolding studied at atomic resolution by fast two-dimensional NMR spectroscopy, Proc. Natl. Acad. Sci. U. S. A. 104 (2007) 11257–11262. [83] E. Rennella, T. Cutuil, P. Schanda, I. Ayala, V. Forge, B. Brutscher, Real-time NMR characterization of structure and dynamics in a transiently populated protein folding intermediate, J. Am. Chem. Soc. 134 (2012) 8066–8069. [84] P. Schanda, H. Van Melckebeke, B. Brutscher, Speeding up three-dimensional protein NMR experiments to a few minutes, J. Am. Chem. Soc. 128 (2006) 9042–9043. [85] D. Lee, C. Hilty, G. Wider, K. Wüthrich, Effective rotational correlation times of proteins from NMR relaxation interference, J. Magn. Reson. 178 (2006) 72–76. [86] B. Fierz, K. Joder, F. Krieger, T. Kiefhaber, Using triplet-triplet energy transfer to measure conformational dynamics in polypeptide chains, Methods Mol. Biol. 350 (2007) 169–187. [87] M. Gruebele, Fast relaxation methods, in: J. Buchner, T. Kiefhaber (Eds.),Handbook of Protein Folding, vol. 1, Wiley-VCH, Weinheim, 2004, pp. 454–490. [88] A. Dose, S. Liokatis, F.X. Theillet, P. Selenko, D. Schwarzer, NMR profiling of histone deacetylase and acetyl-transferase activities in real time, ACS Chem. Biol. 6 (2011) 419–424. [89] B. Fürtig, J. Buck, V. Manoharan, W. Bermel, A. Jaschke, P. Wenter, S. Pitsch, H. Schwalbe, Time-resolved NMR studies of RNA folding, Biopolymers 86 (2007) 360–383. [90] P. Guyett, J. Glushka, X. Gu, M. Bar-Peled, Real-time NMR monitoring of intermediates and labile products of the bifunctional enzyme UDP-apiose/UDP-xylose synthase, Carbohydr. Res. 344 (2009) 1072–1078. [91] I. Amata, M. Maffei, A. Igea, M. Gay, M. Vilaseca, A.R. Nebreda, M. Pons, Multiphosphorylation of the intrinsically disordered unique domain of c-Src studied by in-cell and real-time NMR spectroscopy, ChemBioChem 14 (2013) 1820–1827.

[92] H.J. Dyson, P.E. Wright, Insights into protein folding from NMR, Annu. Rev. Phys. Chem. 47 (1996) 369–395. [93] L.T. Kuhn, Photo-CIDNP NMR spectroscopy of amino acids and proteins, Top. Curr. Chem. 338 (2013) 229–300. [94] K.H. Mok, P.J. Hore, Photo-CIDNP NMR methods for studying protein folding, Methods 34 (2004) 75–87. [95] E. Rennella, B. Brutscher, Fast real-time NMR methods for characterizing shortlived molecular states, ChemPhysChem 14 (2013) 3059–3070. [96] J. Balbach, V. Forge, W.S. Lau, N.A. van Nuland, K. Brew, C.M. Dobson, Protein folding monitored at individual residues during a two-dimensional NMR experiment, Science 274 (1996) 1161–1163. [97] J. Balbach, C. Steegborn, T. Schindler, F.X. Schmid, A protein folding intermediate of ribonuclease T1 characterized at high resolution by 1D and 2D real-time NMR spectroscopy, J. Mol. Biol. 285 (1999) 829–842. [98] T. Kiefhaber, R. Quaas, U. Hahn, F.X. Schmid, Folding of ribonuclease T1. 2. Kinetic models for the folding and unfolding reactions, Biochemistry 29 (1990) 3061–3070. [99] C. Steegborn, H. Schneider-Hassloff, M. Zeeb, J. Balbach, Cooperativity of a protein folding reaction probed at multiple chain positions by real-time 2D NMR spectroscopy, Biochemistry 39 (2000) 7910–7919. [100] C. Löw, U. Weininger, M. Zeeb, W. Zhang, E.D. Laue, F.X. Schmid, J. Balbach, Folding mechanism of an ankyrin repeat protein: scaffold and active site formation of human CDK inhibitor p19(INK4d), J. Mol. Biol. 373 (2007) 219–231. [101] E. Rennella, Z. Solyom, B. Brutscher, Measuring hydrogen exchange in proteins by selective water saturation in H- N SOFAST/BEST-type experiments: advantages and limitations, J. Biomol. NMR 60 (2014) 99–107. [102] W. Doster, J. Friedrich, Pressure-Temperature Phase Diagrams of Proteins, in: J. Buchner, T. Kiefhaber (Eds.),Handbook of Protein Folding, vol. 1, Wiley-VCH, Weinheim, 2004, pp. 99–126. [103] N. Kachel, W. Kremer, R. Zahn, H.R. Kalbitzer, Observation of intermediate states of the human prion protein by high pressure NMR spectroscopy, BMC Struct. Biol. 6 (2006) 16. [104] C.E. Munte, M. Beck Erlach, W. Kremer, J. Koehler, H.R. Kalbitzer, Distinct conformational states of the Alzheimer beta-amyloid peptide can be detected by highpressure NMR spectroscopy, Angew. Chem. Int. Ed. Engl. 52 (2013) 8943–8947. [105] K. Akasaka, Probing conformational fluctuation of proteins by pressure perturbation, Chem. Rev. 106 (2006) 1814–1835. [106] U. Heuert, M. Krumova, G. Hempel, M. Schiewek, A. Blume, NMR probe for pressure-jump experiments up to 250 bars and 3 ms jump time, Rev. Sci. Instrum. 81 (2010) 105102. [107] W. Kremer, M. Arnold, C.E. Munte, R. Hartl, M.B. Erlach, J. Koehler, A. Meier, H.R. Kalbitzer, Pulsed pressure perturbations, an extra dimension in NMR spectroscopy of proteins, J. Am. Chem. Soc. 133 (2011) 13646–13651. [108] C. Schiene-Fischer, T. Aumuller, G. Fischer, Peptide bond cis/trans isomerases: a biocatalysis perspective of conformational dynamics in proteins, Top. Curr. Chem. 328 (2013) 35–67. [109] C. Löw, M.T. Stubbs, C. Haupt, J. Balbach, Metallochaperone SlyD, Encyclopedia of Inorganic and Bioinorganic Chemistry, in: A. Scott (Ed.), John Wiley & Sons, Sussex, 2012, http://dx.doi.org/10.1002/9781119951438.eibc2061 (p.). [110] M. Kovermann, F.X. Schmid, J. Balbach, Molecular function of the prolyl cis/trans isomerase and metallochaperone SlyD, Biol. Chem. 394 (2013) 965–975. [111] R.P. Jakob, G. Zoldak, T. Aumüller, F.X. Schmid, Chaperone domains convert prolyl isomerases into generic catalysts of protein folding, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 20282–20287.

Please cite this article as: A. Kumar, J. Balbach, Real-time protein NMR spectroscopy and investigation of assisted protein folding, Biochim. Biophys. Acta (2014), http://dx.doi.org/10.1016/j.bbagen.2014.12.003