Hydrophobic Collapse of Ubiquitin Generates Rapid ProteinâWater

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Hydrophobic Collapse of Ubiquitin Generates Rapid Protein-Water Motions Hanna Wirtz, Sarah Schaefer, Claudius Hoberg, Korey M. Reid, David M. Leitner, and Martina Havenith Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b00235 • Publication Date (Web): 23 May 2018 Downloaded from http://pubs.acs.org on May 23, 2018

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Biochemistry

Hydrophobic Collapse of Ubiquitin Generates Rapid Protein-Water Motions Hanna Wirtz1, Sarah Schäfer1, Claudius Hoberg1, Korey M. Reid2, David M. Leitner2, Martina Havenith*,1 1 2

Lehrstuhl für Physikalische Chemie II, Ruhr Universität Bochum, 44801 Bochum, Germany Department of Chemistry, University of Nevada, Reno, NV 89557 USA

Corresponding Author *

Lehrstuhl für Physikalische Chemie II, Ruhr Universität Bochum, 44801 Bochum, Germany Telephone: +49 234/32-24249. E-mail: [email protected]

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Abstract We report time resolved measurements of the coupled protein-water modes of solvated ubiquitin during protein folding. Kinetic terahertz absorption (KITA) spectroscopy serves as a label-free technique to monitor large-scale conformational changes and folding of proteins subsequent to a sudden T-jump. We report here KITA measurements at an unprecedented time resolution of 500 ns, two orders of magnitude better resolution than any previous KITA measurements, which reveal the coupled ubiquitin-solvent dynamics even in the initial phase of hydrophobic collapse. Complementary equilibrium experiments and molecular simulations of ubiquitin solutions are carried out to clarify non-equilibrium contributions and reveal the molecular picture upon structural change, respectively. Based upon our results we propose that in the case of ubiquitin a rapid (< 500 ns) initial phase of the hydrophobic collapse from the elongated protein to a molten globule structure precedes secondary structure formation. We find that these very first steps, including large amplitude changes within the unfolded manifold, are accompanied by a rapid (< 500 ns) pronounced change of the coupled protein-solvent response. The KITA response upon secondary structure formation exhibits an opposite sign, which indicates a distinct effect on the solvent exposed surface.


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Introduction Elucidating folding and unfolding mechanisms of proteins remains a central topic in biophysical chemistry.1,2 Many different techniques such as infrared spectroscopy3, time resolved circular dichroism (CD) spectroscopy,4,5 fluorescence spectroscopy,6 and nuclear magnetic resonance (NMR)7 have enabled researchers to gain insight into the mystery of how proteins fold into their native structure. Protein folding and unfolding is accompanied by large-amplitude low frequency motions of the protein, which mediate conformational flexibility and structural dynamics central to protein folding and function.8 Collective protein motions occur on the picosecond (ps) timescale and can be probed spectroscopically in the Terahertz (THz) regime.8,9,10 We could show that the collective modes of the protein are coupled to the collective hydration water network.11–13 When studying the dynamics of water by IR correlation spectroscopy under equilibrium conditions, Groot et al., showed that protein in the first stage unfolding starts with a weakening of protein structure allowing for water uptake.14 Here, we use kinetic THz absorption (KITA) spectroscopy as a label-free method to track folding and unfolding of proteins.15,16 Here, we want to address how the collective protein – water motions are affected upon protein folding under non-equilibrium conditions. Protein folding is characterized by a hydrophobic collapse as well as secondary structure formation. These steps might proceed sequentially or in parallel. Even less is experimentally known about the very first steps of the collapse starting from elongated protein structures up to a well-defined structure. Femtosecond THz absorption spectroscopy allows us to monitor the changes in collective protein hydration dynamics even at times of less than 500 ns after the initiation of protein folding. Here we report time-resolved THz spectroscopy on a sub-µs time scale to monitor, via the THz response, rapid protein structural changes following a rapid temperature jump initiating folding of a cold denatured protein. Most of the studies on solute-water dynamics are carried out in the steady-state, i.e. under equilibrium conditions. Studies were reported for human serum albumin (HSA)11, lambda repressor (6-85-repressor)13,17, ubiquitin (UBQ)13,18 or lysozyme.19,20 Our group has developed KITA (kinetic THz absorption) spectroscopy to study coupled collective protein-water dynamics by means of THz absorption spectroscopy ACS Paragon Plus Environment

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in combination with a kinetic stopped-flow apparatus.15 Rapid mixing of the denatured protein solution (pseudo-wild-type fluorescent Phe45Trp mutant of UBQ) with buffer in a stopped-flow mixer induced the folding of the protein. The change in THz amplitude was recorded as a function of delay time after refolding was initiated. Due to the dead time of the stopped-flow mixer, however, the time resolution was restricted to 1 ms. We successfully demonstrated that KITA spectroscopy is sensitive to changes in the protein-water dynamics during folding and found that the hydrophobic collapse is correlated with a considerable change in the solvent dynamics that precedes final tertiary structure formation.15,17 We assigned the relaxation kinetics to intermolecular hydrogen bond formation in early events of protein folding and gave an upper limit of 11 ms for the relaxation time. However, due to the relatively large dead time of the stopped-flow mixer, it was not possible to timeresolve the kinetics. Conformational changes and enzymatic catalysis were found to be correlated with protein-water coupled motions.21 Protein or enzymatic function is drastically reduced in the absence of water, thus indicating that water plays a significant role in biomolecular and cellular processes.22,23 Using KITA spectroscopy, we investigated the human membrane type-1 MMP (MT1-MMP) and recorded the changes in the protein-water coupled motions during peptide hydrolysis by a zinc-dependent human metalloprotease.21,24 Our results indicated that these changes were strongly correlated with structural rearrangements at the active site during the formation of enzyme-substrate intermediates.21,24 Recently, we combined KITA with a fast nanosecond temperature jump, thereby improving the time resolution to 50 µs.16 For 6-85-repressor we found a slow relaxation on the time scale of ms, which matched the time constant of the hydrophobic collapse. Here, we report KITA studies on UBQ, which is an ideal model system to investigate protein structure, stability, folding and dynamics.25 The protein is ubiquitous in cellular environments and consists of 76 amino acids (radius of gyration of 11.7 Å).13 It has been shown that UBQ is extraordinarily stable against temperature (23 to 80 °C) and pH (1.2 to 8.5) changes as well as denaturant concentrations. At pH 2, UBQ undergoes heat denaturation at a melting temperature of 57 °C.26 Wand and coworkers made use of reverse micelles to study the cold denaturation transition of


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Biochemistry

UBQ between -20 and -30 °C and showed that unfolding is highly non-cooperative.7 Ibarra-Molero, et al., used differential scanning calorimetry (DSC) measurements and found that the heat denaturation transition could be shifted to lower temperatures when increasing the concentration of GdmCl from 0 to 2 M.27

Unlike 6-85-repressor, in UBQ folding the hydrophobic collapse is postulated to precede secondary structure arrangement.28–32 Fluorescence measurements of the folding of a UBQ mutant (F45W V26G) revealed kinetics with a fast 22 µs and a slow 2 - 5 ms phase, the latter being attributed to native structure formation.33 Studies on the native-like F45W mutant of UBQ provided evidence that an early folding intermediate is populated during the first few ms of refolding.32 Roder, et al., observed a 50 % change of the fluorescence signal within the 2 ms dead time of the instrument.32 However, on a time scale of 5 ms and beyond a 73 % change could be observed. Kinetic CD spectra indicate a single phase of 10 ± 3 ms for the refolding of UBQ from a guanidine hydrochloride (GdmCl) denatured state at 20 °C.34 Briggs and Roder used hydrogen/deuterium exchange labeling in combination with 2D NMR to investigate the early events of the UBQ folding reaction. They could distinguish three time constants: 8 ms, 100 ms and 10 s.30 In the initial fast 8 ms folding phase 80 % of the amide protons in secondary structure elements and protons involved in hydrogen bonding between these elements become protected, thus indicating that most of these elements form and associate in a common and cooperative folding event. Based upon all-atom molecular dynamics simulations from Marianayagam and Jackson it was proposed that folding of wild type UBQ is initiated by a hydrophobic collapse phase followed by subsequent major secondary structure rearrangement.28 They concluded that the initial collapse of UBQ is most likely not observable with the majority of experimental techniques due to the large dead times of the instruments (ms time scale). In the present study, we investigate the T-jump initiated folding/unfolding transition of the protein UBQ from bovine erythrocytes at low temperatures with a time resolution of 500 ns. By means of KITA measurements it is possible to detect changes in water network dynamics caused by the structural reorganization of the protein during the folding transition. Accompanying equilibrium experiments and molecular simulations


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clarify the contributions due to structural changes and thermal effects on the KITA measurements.

Materials and Methods Sample Preparation Ubiquitin from bovine erythrocytes (Mw= 8565 Da, ≥ 98 % purity) was purchased from Sigma

Aldrich

(Germany)

and

used

without

further

purification.

For

all

measurements, the protein was dissolved in a pH 2 (pH meter, Hanna Instruments, Italy) buffer solution containing 10 mM glycine (≥ 99.0 % purity, Sigma Adrich, Germany) and 3.55 M GdmCl (≥ 99.5 % purity, Sigma Adrich, Germany). The pH of the buffer was set with hydrochloric acid (VWR International GmbH, Germany). Both static and kinetic THz measurements were carried out using a concentration of 5 mg/mL.

CD spectroscopy CD measurements were performed using a conventional spectrometer (JASCO J715, United States). The sample was measured in a quartz cuvette with a cell path length of 0.1 mm. Data were collected for temperatures ranging from -12.5 to 20 °C using a thermostat (Julabo F25, Germany). The exact concentration of the measured protein sample was determined via UV/Vis spectroscopy (NanoVue 4282 V2, Biochrom, Germany) at 280 nm using an extinction coefficient for UBQ of 0.149 mg1

cm-1Ml yielding a concentration of 1.8 mg/mL.35

THz Time Domain Spectrometer THz absorption measurements of the solutions in equilibrium were carried out using a THz time-domain (THz TD) spectrometer. A detailed description of a THz TD spectrometer can be found in [16, 36, 37]. In our custom-built setup, a Ti:Sa laser (MaiTai DeepSee eHP SpectraPhysics, United States) emits mode-locked 800 nm pulses at an 80 MHz repetition rate with a pulse duration of 70 fs and an average output power of 3 W. The 800 nm pulse is split into a pump (96%) and a probe beam (4%) by a beam splitter. The pump pulse propagates via a variable delay line and is focused on a GaAs photoconductive THz antenna (TeraSED3, GigaOptics,


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Biochemistry

Germany) biased and modulated with 20 V and 92 kHz, respectively. The emitted THz field (0.2 - 3 THz) is focused onto the sample via two off-axis parabolic mirrors. The sample cell is composed of two z-cut quartz windows separated by a 100 µm thin teflon spacer. After passing through the sample, the transmitted pulse is focused on a 0.5 mm (110)-ZnTe crystal by two additional off-axis parabolic mirrors and overlapped with the probe pulse for electro-optical detection. The electric field of the THz pulses induces birefringence in the ZnTe crystal. This causes a change in polarization, which is proportional to the electric field of the THz pulse. The net change is detected by a balanced photodetector (125 kHz, Nirvana Auto-Balanced Photoreceiver Model 2007, New Focus, United States) probing both polarization states and processed by a Lock-in amplifier (HF2-LI, Zurich Instruments, Switzerland). Using a fast delay stage (2 Hz, ScanDelay 150, APE, Germany) in the pump beam path it is possible to observe the THz pulse in real time covering a time window of about 17 ps. For static THz measurements, a time constant of 1 µs with a filter of 3 dB was used. Using these parameters, a dynamic range of 50 dB was achieved after 30 s of integration for water measurements. Temperature was controlled by an external thermostat with 0.1 °C accuracy.

T-jump Setup The setup used for KITA measurements is shown in Fig. 1. A Q-switched Nd:YAG laser pulse (1064 nm, Surelite III-10, Continuum, United States) with an output power of 800 mJ/pulse and a pulse duration of 5 ns is focused into a methane gas Raman shifter (1500 psi, 101 PAL, Light Age, United States). By stimulated Raman emission, the incoming 1064 nm laser pulse is converted to 1540 nm, a wavelength close to water’s overtone absorption.36 With an average conversion efficiency of 15 %, a pulse energy of 120 mW at 1540 nm is gained after the Raman shifter. To enable smooth heating throughout the solution, a double pump configuration from two sides is used. The pulse is split into two parts by a beam splitter with high damage threshold and independently focused onto the sample cell, where they overlap in space with the THz pulse. At the sample position, both heating pulses have larger beam waist (2 mm) than the THz pulse (~ 700 µm), thus probing of only the heated volume is guaranteed.


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Figure 1: Schematic of the KITA Spectrometer in double-pump configuration.

The THz emitter is modulated with a frequency of 2 MHz and THz absorption is detected by a 80 MHz balanced photodetector. Processing and digitization occurs via a high-speed AD converter (Octopus 2, GaGe Applied Technologies Inc.) with a sampling rate of 125 MHz. For KITA measurements, a Nd:YAG repetition rate of 2 Hz was chosen to enable thermal relaxation to equilibrium conditions controlled by an external constant bath. Data sets with a time window of 25 ms (6.25 ms before the Tjump) are recorded for 1.5h (10800 heating traces) and averaged. Time resolution is limited by the antenna modulation frequency of 2 MHz resulting in a time resolution of 0.5 µs. For all measurements, a background subtraction (signals detected without THz light emitted) is carried out to remove reflection artifacts from T-jump traces.

Molecular Simulations Using the GROMACS molecular dynamics package37 and the CHARMM36m force field for the protein, an initial structure (PDB: 1UBQ) was solvated with the TIP3P water model in a triclinic box with a solvation buffer radius of 1.2 nm and NaCl concentration of 0.15 M. The box size was sufficient to accommodate the protein, either in its folded or unfolded structure. To provide a comparison to the solvation water dynamics we simulated a triclinic water box of equal size, side lengths of 6.89 nm, filled with TIP3P water to simulate bulk water. The simulations were integrated using a 1 fs time-step with the Verlet integrator, a cutoff of 1.4 Å was employed for


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Coulomb and van der Waals forces, electrostatic forces were evaluated using particle mesh Ewald and LINCS was employed to constrain bonds including hydrogen. To generate unfolded structures, we took the starting ubiquitin system and unfolded the structure at two different temperatures, 700 K and 1000 K. Each was heated under an isobaric-isothermal ensemble (NPT) at a rate of 1 K ps-1 and position restraints applied to heavy atoms with a force constant of 1000 kJ mol-1 nm-2. Upon reaching 300 K the pressure was increased to 5 kbar, to more efficiently unfold the protein, and heating continued without restraints until the final temperatures were reached. Each system was allowed to equilibrate for an additional 1.2 ns producing the final unfolded starting structures. For all three structures two simulations were conducted in succession, first NPT for 1 ns then NVT for 300 ps at 300 K. These simulations were carried out again in a triclinic box, with a solvation buffer radius of 1.2 nm, and NaCl concentration of 0.15 M. The starting structures for normal mode analysis were taken from the final structure of the NPT simulation and vibrational density of states (VDOS) power spectra were calculated using the NVT simulation. The final structures of the NPT production

runs

were

energy

minimized

for

normal

mode

analysis.

The

eigenfrequencies, eigenvectors and Hessian matrix were used in the calculation of thermal transport coefficients of the proteins.38 To obtain the vibrational density of states (VDOS), the velocity autocorrelation function (ACF) for the oxygen atoms of the water molecules was calculated in the final 100 ps of the NVT simulation at 300 K, evaluated using a 15 ps ACF length with a time step of 5 fs. All water molecules were initially within a 5 Å shell around the protein and only those remaining in the hydration layer for the duration of the 100 ps simulation were evaluated, for a total of about 70 water molecules around the protein.

Results Cold denaturation of Ubiquitin We carried out CD measurements in the temperature range from -12.5 to 20 °C. The CD spectra show two prominent signals at 222 and 208 nm, which indicates the ACS Paragon Plus Environment

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Figure 2: CD signal at 222 nm plotted against the temperature.

presence of -helices in the protein.39 With decreasing temperature, the signal at 222 nm increases, thus indicating a transition from the folded protein with intact -helices to a random coil conformation.40 In order to determine the temperature for cold denaturation, the CD signal at 222 nm is plotted against the respective temperature (Fig. 2). The data was fit using the following form of a Boltzmann function,

ΘMRW = A2 +

A1 + A2  T − Tm  1+ exp    dT 

(1)

where Θ is the mean residue ellipticity, A1 and A2 are the initial and final value for Θ , Tm is the melting temperature, and dT is the temperature constant. As a result we obtain A1 = 0.30 ± 0.19, A2 = -3.92 ± 0.15, Tm = (1.81 ± 0.63) °C and dT = (5.86 ± 0.74) °C. The melting temperature upon cold denaturation (1.81 ± 0.63) °C agrees well with the extrapolated data from Ibarra-Molero.27,35

Based on the results from the Boltzmann fit, 100 % unfolded structure (U) accounts for a CD signal or an amplitude A1 of 0.30 ± 0.19, whereas 100 % folded structure (F) corresponds to a CD signal of A2 = -3.92 ± 0.15.

Equilibrium THz measurements It is well-known from former studies that the absorption of water increases linearly with increasing temperature for frequencies ranging from 0.1 to 2 THz.41 In this study, equilibrium measurements were carried out using our THz-TD spectrometer (0.2 – 3 THz) for the buffer (10 mM glycine, pH 2, with/without 3.55 M GdmCl) and protein ACS Paragon Plus Environment

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Biochemistry

solutions (5 mg/ml dissolved in respective buffer). Cell temperature was controlled within the range of about - 0.5 °C to + 20 °C, thereby covering the transition temperature for cold denaturation. The transmitted electric field of the THz pulse was detected subsequently for an empty (reference) and a filled cell (sample) and the frequency-dependent absorption coefficient was deduced.

All changes follow a linear behavior. The respective slopes of the linear fits are displayed in Table 1. The addition of GdmCl on the absorption coefficient causes an increase in absorption. The increase shows a linear dependence in temperature, similar as for the buffer itself (see SI).

Table 1: Decrease of transmitted THz amplitude of the solution with increasing temperature Solution UBQ without GdmCl Buffer without GdmCl UBQ with GdmCl Buffer with GdmCl

Fitted slope (/°C) 0.0194 ± 0.0007 0.0169 ± 0.0010 0.0135 ± 0.0007 0.0112 ± 0.0003

In Fig. 3 we show the difference in absorption ∆ for distinct temperatures and concentrations. ∆ was calculated according to ∆ / within three distinct frequency ranges. Note that ∆ is not density corrected. However, the protein concentration is very low (0.58 mM), thus we expect no influence of this assumption within experimental error.

Figure 3: ∆α for distinct temperatures and concentrations averaged over different frequency regions. The data are calculated averages of three independent measurements.


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We observe a small, yet significant difference in ∆ for the unfolded (0 °C) versus the folded (20 °C) protein, which is more pronounced at higher frequencies (2–2.4 THz). Similar as was found for the protein HSA, the solvated unfolded protein absorbs significantly less THz light than the folded protein.11 This change in THz absorption was attributed to the fact that the solvent exposed surface for the unfolded protein is considerably increased compared to the folded protein.

MD Simulations Using molecular simulations, the vibrational density of states (VDOS) of water molecules around UBQ was calculated for structures with varying radius of gyration (Rg). Two unfolded structures with Rg = 1.5 Å (red) and Rg = 1.7 Å (green) and one folded structure (Rg = 1.2 Å (blue)) were investigated. For comparison, the VDOS of bulk water is shown in black. Figure 4 represents the VDOS for the nearest 70 water molecules, which are about 3 Å away from the surface.

Figure 4: Calculated VDOS for the nearest 70 water molecules around unfolded (red: Rg 1.5 nm, green: 1.7 nm) and folded (blue Rg 1.2 nm) UBQ. The VDOS for bulk water is shown in black for comparison.

In the presence of UBQ the VDOS for hydration water is blue-shifted and the amplitude decreases compared to bulk water. This trend is consistent with previous studies of water around proteins, where the absorption coefficient of water around 1 THz is found to be decreased in the presence of a protein.13,16,42 At low frequency such a shift is expected, since the 0-frequency VDOS is proportional to the diffusion coefficient, which is larger for water in the bulk than near the surface of a protein, ACS Paragon Plus Environment

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Biochemistry

where the dynamics is more retarded. That dynamic can also be sensitive to protein structure, as is evident in Fig. 4, where there are noticeable differences in the VDOS at low frequency.

For the three investigated structures, the native (1.2 nm) and the two unfolded (1.5 and 1.7 nm) structures, the VDOS is slightly higher for the two unfolded structures than for the native one. We expect that the VDOS will be blue shifted at low frequency for the unfolded structures, since the 0-frequency VDOS is proportional to the diffusion coefficient. The observed blue shift at low frequency is consistent with greater mobility of the water near the protein when it is unfolded, the result of more exposure to hydrophobic residues. Interestingly, it is slightly greater for the structure with a radius of gyration of 1.5 nm than 1.7 nm. Though we generally expect a blue shift as the protein unfolds, the extent of the shift will vary for unfolded structures since the mobility of the water around the protein depends on contacts with exposed residues and on the surface structure. This

choice

of

structures

was

motivated

by

time-resolved

fluorescence

measurements indicating rapid hydrophobic collapse of UBQ to a molten globule state with a radius of gyration that is comparable to 1.5 nm,31 so that for collapse into the molten globule state the VDOS may increase. Of course, we do not know how representative the selected unfolded structures used to obtain the VDOS are of the ensemble of molten globule structures, or for the ensemble of structures under equilibrium conditions. However, we can conclude that the VDOS of water around UBQ is sensitive to structural change and is in general lower than the VDOS for bulk water, which is expected, as discussed above. This is in agreement with the experimental observation of a decrease in the response of solvated protein solution compared to bulk water under equilibrium conditions. In addition, thermal diffusion coefficients were calculated for solutions of UBQ in folded and unfolded structures. The thermal diffusivity for the bare protein in its folded state is found to be smaller (0.1 mm²/s) than when unfolded (0.2 mm²/s) at 20 °C. Accounting for the protein concentrations used in the experiments the thermal diffusivity of the protein solutions remains within 0.15% of its value for water (0.143 mm2/s) when the protein is either in a folded or unfolded structure.


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Kinetic Terahertz Absorption Spectroscopy upon T-jump We use temperature jumps with a time resolution of 500 ns to initiate the refolding of UBQ and follow the THz absorption starting from a cold denatured protein. T-jump traces are recorded at the maximum peak position of the THz pulse for 6.25 ms before and 18.75 ms after the T-jump. The THz amplitude before the T-

! with n representing the total number of data jump (t < 0 ms) is averaged to points before the jump (12,500 points). The KITA signal is then calculated as "#$

∆%&'( )*+, - %!&'( )*+,

%&'( )*+, -

∑0 12 3.56 12 %&'( )*+, - //

(2)

Prior to the T-jump, the protein solutions were equilibrated at a temperature of -1 °C. When a T-jump is applied, the solution is instantly (within < 500 ns) heated up by 6 – 7 °C, initiating protein refolding. Each data point in Figure 5 represents the average over 15.000 laser shots of the T-jump laser (NdYAG), thus signal shot power fluctuations will cancel. According to the results of CD spectroscopy, such a T-jump should induce a change in the equilibrium (U ↔ N) of about 25 %. Fig. 5 shows that the THz amplitude decreases rapidly for both buffer and UBQ. For the buffer solution,

Figure 5: Normalized T-jump trace for UBQ and buffer. Shown is the change in THz Amplitude at time t=0 ps. Inset: zoom-in into the sub-millisecond regime.

the change in amplitude amounts to a decrease in ETHz of 7.2 %, whereas for UBQ solutions a decrease of 7.5 % is recorded.


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Biochemistry

Subsequent to the T-jump, the solution cools down and relaxes to equilibrium conditions within ~50 ms. The cooling rate is diffusion limited and the relaxation can be described by 16

y = y0 + A1e−t/τ1 + A2 e−t/τ 2 .

(3)

The result of the fit is plotted in Fig. 5.

τ1 and τ2 have contributions from the diffusion

limited cooling process as well as protein folding.

Table 2: Results for a bi-exponential fit to the data in Fig. 5 parameter

Ubiquitin

Buffer

τ1 (ms)

0.847 ± 0.036

0.715 ± 0.033

τ2 (ms)

9.657 ± 0.088

9.383 ± 0.076

Α1

0.00878 ± 0.00018

0.00770 ± 0.00017

Α2

0.053882 ± 0.000095

0.049974 ± 0.000084

In Fig. 6, the difference of the two T-jump traces is plotted against time. We observe a negative difference ∆KITA signal when subtracting the KITA signal of the buffer+GdmCl from the protein signal: KITAProtein – KITABuffer. The difference signal (for t > 0 ms) can be fitted to a single exponential with a time constant of (7.546 ± 0.060) ms.

Figure 6 Difference in KITA Signal. The buffer+GdmCl trace was subtracted from the ubiquitin trace. A time constant of (7.546 ± 0.060) ms. was obtained using a single exponential fit.


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We propose that the KITA signal is correlated with the change of the collective hydrogen bond network dynamics during protein folding. As control measurements, we have recorded the change in THz absorption upon T-jump at 52 °C and -1 °C for a solution of UBQ with 10 mM glycine at pH 2. Under these conditions, UBQ is expected to be stable.26,27,35

Figure 7: Normalized KITA signal for UBQ and buffer (without GdmCl) for a T-jump starting at 52 °C (left) and -1 °C (right). The insets show the difference in KITA signal.

We find no difference in the response for ubiquitin and buffer within experimental uncertainty (Fig. 7), which confirms our initial assumption.

Discussion First of all, we have to exclude the possibility that the difference in the KITA signal can be caused by changes in thermal diffusivity or ∆89 : the thermal diffusivity of water, 0.143 mm2/s at 20 ˚C, is affected by the presence of protein. However, for the concentrations used in the KITA measurements, any differences of thermal diffusion in buffer and protein solutions is too small to cause measurable changes in the absorption coefficient between the two solutions. Furthermore, protein solution and buffer exhibited the same time-dependent absorbance changes for any case where the protein structure is maintained. Following a T-jump, UBQ is postulated to collapse to an intermediate molten globule state, which is accompanied by a sudden change in heat capacity. The change in heat capacity as deduced by calorimetry

26,43

is ca. 0.00021 cal/K for the unfolded


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state and 0.000138 cal/K in the native form at 5 °C for a cell thickness of 100 µm. Based on the assumption that the entire energy of the heating pulse is absorbed, this will yield a temperature jump of ∆ = 8.42 K for a buffer solution containing 3.55 M GdmCl at 10 °C (89,;. 0 ms reflects the change in the coupled protein hydration water during partial secondary structure formation while unfolding starts, also in agreement with the predicted VDOS. In summary, we find that KITA is a very sensitive label-free probe of the collective large amplitude changes during protein conformational changes. As shown in the present study, real time tracking of protein folding by KITA is possible. It provides complimentary results to real time tracking of secondary structure formation (as provided by fluorescence or CD spectroscopy). Even at protein concentrations of 1 in roughly 104 water molecules, the change in the coupled water-protein dynamics is large enough to be detected, which indicates a remarkable imprinting on the water network. The sign of the change with respect to buffer yields information on the protein folding process even in the very first phase of protein folding: while ∆KITA is negative in the case of an initial rapid hydrophobic collapse from an elongated protein to a molten globule (ubiquitin), the initial response for protein folding preceding via secondary structure formation followed by slow hydrophobic collapse ( 6-85-repressor) yields a positive ∆KITA.16 Our study shows that water responds closely to any structural changes. Each step in this process is accompanied by clearly detectable solvation changes. The sub-µs time resolution achieved here, two orders of magnitude faster than the resolution of previous KITA measurements, allows us not only to monitor the solvent response during hydrophobic collapse of a protein preceding secondary structure formation, but also to directly observe the changes while the collapse is still taking place.

Supporting Information Full CD-spectra of UBQ, Temperature dependence of THz-pulses, Equilibrium THz spectra of UBQ and buffer, Explanation of the relation between ∆T and ∆ETHz , ∆KITA signal for buffer with and without GdmCl, simulated structures of native and unfolded UBQ (Rg = 1.7 nm and Rg = 1.5 nm). ACS Paragon Plus Environment

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Acknowledgment We thank Janne Savolainen for initial help with the set-up of the experiment. This work is part of the Cluster of Excellence Ruhr Explores Solvation (RESOLV) (EXC 1069) funded by the Deutsche Forschungsgemeinschaft. Additional funding was provided by NSF CHE-1361776 (DML).


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