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Dynamics of hydration water and coupled protein sidechains around a polymerase protein surface
Chemical Physics Letters xxx (2017) xxx–xxx
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Dynamics of hydration water and coupled protein sidechains around a polymerase protein surface Yangzhong Qin, Yi Yang, Lijuan Wang, Dongping Zhong ⇑ Department of Physics, Department of Chemistry and Biochemistry, Programs of Biophysics, Chemical Physics, and Biochemistry, The Ohio State University, Columbus, OH 43210, United States
a r t i c l e
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Article history: Received 31 December 2016 In final form 2 March 2017 Available online xxxx Keywords: Hydration dynamics Slaved sidechain motions Water-protein coupled fluctuations Femtosecond spectroscopy Tryptophan scan
a b s t r a c t Water-protein coupled interactions are essential to the protein structural stability, flexibility and dynamic functions. The ultimate effects of the hydration dynamics on the protein fluctuations remain substantially unexplored. Here, we investigated the dynamics of both hydration water and protein sidechains at 13 different sites around the polymerase b protein surface using a tryptophan scan with femtosecond spectroscopy. Three types of hydration-water relaxations and two types of protein sidechain motions were determined, reflecting a highly dynamic water-protein interactions fluctuating on the picosecond time scales. The hydration-water dynamics dominate the coupled interactions with higher flexibility. Ó 2017 Elsevier B.V. All rights reserved.
1. Introduction Water is a necessity for nearly all life known [1]. It involves in most biological processes not only as a universal solvent, but also as an active participant in mediating protein folding [2–4], protein-ligand binding [5–7] and enzymatic catalysis [8–10]. Hydration water, directly interacting with protein at the waterprotein interface, strongly influences the protein structure and dynamics [11–15]. Mutually, the water structure and dynamics are significantly perturbed as well [16–21]. Understanding the dynamics of coupled water-protein interactions is crucial to revealing its ultimate effects on the protein’s functions that require both local and global flexibility in aqueous solution. Extensive studies had been carried out to investigate the dynamics of either hydration water or proteins by both experiments [22–28] and simulation [14,20,21,29]. However, due to the strong water-protein coupling, it remains challenging to disentangle their dynamics. Here, we report our simultaneous measurements of the surface hydration dynamics and related protein sidechain motions by employing the femtosecond (fs)-resolved fluorescence spectroscopy and a tryptophan (W) scan [18,19]. With site-directed mutagenesis, we placed a single tryptophan, one at a time, as an optical probe at 13 different locations on the surface of DNA polymerase b (Pol b) to globally map out the dynamics of hydration water and protein sidechains (Fig. 1). The structure and function ⇑ Corresponding author. E-mail address: [email protected] (D. Zhong).
of Pol b have been extensively studied previously [30–34]. As an X-family DNA polymerase, it can bind to the gapped or nicked DNA and fulfill gap filling synthesis. Pol b has 335 amino acids (39 kDa) with the overall structure following the right hand architecture (Fig. 1). The polymerase domain (31 kDa) consists of three subdomains: a double-stranded DNA (dsDNA) binding subdomain D (also called thumb), a nucleotidyl-transfer subdomain C (palm), and a dNTP selection subdomain N (finger). The N-terminal lyase domain (8 kDa) performs the lyase function to remove the 50 -deoxyribose phosphate during base excision repair [32]. All tryptophan probes were scattered on different domains of Pol b to maximally sample the protein surface heterogeneity. Meanwhile, all tryptophan probes except W325 (WT) are designed at or close to the DNA binding sites that can examine the hydration dynamics of binding sites contrasting to non-binding sites in the apo state [31]. 2. Materials and methods The plasmid of rat pol b (PET17b) was kindly provided by the Tsai’s group at the Ohio State University. Wild type pol b contains a single tryptophan at W325, which was mutated to W325F and served as the double mutant template. Twelve double mutants, each with a single tryptophan, were designed. The plasmids of wild type and all twelve double mutants were overexpressed in Escherichia coli BL21(DE3) one by one. Each protein was then purified following previously reported protocol [34]. We further checked the CD spectra and confirmed the similar structures of the mutants
http://dx.doi.org/10.1016/j.cplett.2017.03.002 0009-2614/Ó 2017 Elsevier B.V. All rights reserved.
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Fig. 1. Ribbon and surface structures of pol b in the apo state (Protein Data Bank ID: 1BPD). Pol b includes a lyase domain (gray) and three polymerase subdomains, thumb (green), palm (red), and finger (blue), forming a right-hand configuration. Yellow spheres and patches indicate the positions of tryptophan probes. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
to that of WT. Each protein was finally stored in a buffer containing 100 mM Tris-HCl (pH 8.0), 150 mM KCl, 2 mM DTT, 3% glycerol (vol/vol), and 1.5 mM MgCl2. The final protein concentration is about 400 lM. The complete details of the time-resolved fluorescence spectroscopy were reported previously [35]. In summary, we used the fluorescence up-conversion technique. An 800 nm pulsed laser beam with 1 kHz repletion rate and 110 fs temporal width was generated by a commercial amplifier (Spitfire, Spectra-Physics). It was split into two parts, with 10% to serve as the probe beam and 90% to pump an optical parametric amplifier (OPA 800C, Spectra-Physics). The OPA generated 590-nm laser pulses that traveled through the translation stage with maximum 3 ns delay relative to the probe. Then, it was focused on a doubling BBO crystal to produce 295 nm pump pulses. Each pump pulse energy was attenuated to 140 nJ before focusing on the protein sample stored in a motor-controlled spinning cell with 1 mm thickness. The protein fluorescence was collected and focused on the up-conversion BBO crystal to mix with a focused probe pulse. The up-converted signal was directed to a monochromator coupled with a photomultiplier tube (PMT). The instrument response time under current non-collinear condition is around 450 fs determined by water Raman signal at 325 nm. For all the solvation transients, the pump pulse polarization was set to the magic angle (54.7°) with respect to the acceptance axis of the up-conversion BBO crystal. For fluorescence anisotropy study, the polarization of pump pulse was set to be either parallel or perpendicular to the acceptance axis to record the parallel signal (I//) and perpendicular signal (I\), respectively. The probe beam polarization was always parallel to the acceptance axis of the up-conversion BBO crystal. 3. Results and discussion 3.1. Femtosecond-resolved fluorescence transient dynamics For each mutant, we have measured the fluorescence transients with a 3 ns time window at different wavelengths from 310 nm to 370 nm covering the blue- and red-side emission spectrum. Fig. 2 shows the typical fluorescence transients of a buried mutant G179W and an exposed mutant S109W. Detailed data-analysis strategy was provided previously [35]. We observed typical solvation patterns for all mutants. For the buried mutant G179W, beside two lifetime decays in nanoseconds (1.2 ns and 5.5 ns), we observed two picosecond solvation decays (4.5–11.2 ps and 82– 108 ps) at the blue side and one rise component (3.8–7.0 ps) at the red side of the emission spectrum. For the exposed mutant S109W, we observed three solvation decays (0.38–0.6 ps, 3.6– 5.0 ps and 34–51 ps) at the blue side and one rise component
Fig. 2. Normalized fluorescence transients at a series of wavelengths for the buried mutant G179W (A) and exposed mutant S109W (B). The scatter plots and the solid lines show the raw data and best multi-exponential fits, respectively.
(0.8–1.1 ps) at the red side in addition to the two lifetime decays (0.7 ns and 5.1 ns). All these decay and rise dynamics are typical solvation signature, which has been observed in many other proteins [18,19], reflecting the relaxations of excited tryptophan and its surroundings. 3.2. Solvent exposure, solvation correlation function and hydration dynamics To further understand the solvation dynamics measured by tryptophan at different locations, we need to know the environ-
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ment of tryptophan probe such as its local protein topology and solvent exposure. Fig. 3A presents a snapshot of MD simulations showing the protein structure as well as hydration water detected by tryptophan for mutant G179W and S109W. The steady-state emission peak can roughly serve as an indicator for the solvent exposure. Fig. 3B shows the steady-state emission spectra of selected four mutants. Bluer emission indicates buried environment of tryptophan and less solvent exposure while redder emission indicates the exposed position of tryptophan and higher solvent exposure [36]. The gradual change of solvent exposure can be approximately quantified by the steady-state emission peaks listed in Table S1 of the supplementary material. Typically, with the dividing line around 338 nm of the emission peak (kpeak) [36], we can separate different mutants into two groups: G179W and F146W as a buried group and all the others as an exposed group. With the fluorescence transients gated at different wavelengths, one can construct the fs-resolved emission spectra (FRES) at different times and monitor the dynamic Stokes shifts with time. As demonstrated in previous studies [18,19], the solvation correlation
Fig. 3. (A) A snapshot of MD simulations shows the water molecules detected within 10 Å from the indole ring of tryptophan for the buried mutant G179W and exposed mutant S109W. Buried tryptophan only detects inner-layer (red) hydration water within 7 Å from the protein surface and exposed tryptophan detects both inner- and outer-layer (blue) hydration water. (B) The steady-state emission spectrum gradually shifts to the red side when the tryptophan is moving from a buried to exposed position, with a dividing line around 338 nm. (C) The solvation correlation functions report either two or three water relaxation processes by the buried mutant G179W and exposed mutant S109W, respectively.
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function C(t) can be deduced, which directly measures the solvation relaxation. Fig. 3C shows the solvation correlation functions for both the buried mutant G179W and exposed mutant S109W. Consistent with fluorescence transients, the solvation correlation function exhibits only two decays (6.7 ps and 106 ps) for the buried mutant and three decays (0.53 ps, 3.9 ps and 47 ps) for the exposed mutant (see Table S1 for details). The ultrafast component detected by the exposed mutant comes from the outer-layer water of the hydration shell that is distant from the protein surface. Indeed, our recent studies in many proteins suggest that buried mutants only detect hydration water within 7 Å of the inner hydration layers from the protein surface [37–39], but exposed mutants can detect water molecules beyond 7 Å forming the outer hydration layers as shown in Fig. 3A. Fig. 4 shows the distributions of solvation energies and solvation relaxation times of all mutants. The total solvation energy (Etot) and the ultrafast solvation energy (E1) gradually increase when the solvent exposure is increasing. The E2 and E3 increase before the emission peaks reach 338 nm and stay nearly no change afterwards. This suggests that both E2 and E3, especially E3, mainly come from the inner-layer hydration water response. It also suggests that the protein solvation contribution may be negligible, consistent with our recent studies [37–39], because both E2 and E3 keep decreasing when solvent exposure reduces. Detailed solvation energies and solvation times are listed in Table S1. The ultrafast solvation happens in sub-picosecond for all exposed mutants, close to the bulk-water relaxation around 0.34 ps [37]. Therefore, it reflects the bulk-type nature of the outer-layer water of the hydration shell. On such an ultrafast time scale, the solvation mainly comes from the water libration and hindered rotation. The second solvation occurs in 3.9–8.7 ps (s2S) during which water molecules typically undergo large amplitude reorientations [40,41]. The slowest relaxation happens in 47–106 picoseconds, reflecting the whole water network reorganizations, coupled with local protein fluctuations. This time scale is consistent with the surface hydration diffusion time and also residence time reported by NMR studies as well as by simulations [24,42,43]. Due to the close interactions of inner-layer hydration water with the protein, the slow relaxation times show clear distinctions that are correlated to the protein surface heterogeneity. For an example, hydration water around hydrophobic sites (S109W and K41W) show relatively faster relaxation times for both water reorientations and water network reorganizations. The heavily charged environment shows the slowest relaxations. Local protein steric confinement also plays an important role in slowing down the hydration-water relaxation. Buried mutants of F146W and G179W basically detect inner-layer hydration water molecules that are confined on the rugged protein surface. Although L22W is categorized as an exposed mutant, the local protein surface forms deep pockets and clefts, rendering high restriction to the inner-layer hydration water and hence causing water molecules to move slower. All these results lead us to the conclusion that the protein surface hydration dynamics are directly coupled to the protein local environments such as hydrophobicity, charge density and steric confinement. This is in agreement with previous studies both experimentally [35] and computationally [41]. Except WT, all other mutants were designed to have the tryptophan probe placed around the DNA binding site. The WT protein serves as a control to report the hydration dynamics at nonbinding sites and all other mutants report the hydration dynamics at the DNA binding sites. As shown in Fig. 4, no clear distinctions were observed for the hydration dynamics at the DNA binding sites compared to that of the non-binding site (WT). However, this is only true for the apo state without binding to DNA. Previous studies on the binary (pol b-DNA) and ternary (pol b-DNA-dNTP) complex states showed a moderate slowdown by a factor of 2 for the
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Fig. 4. Solvation dynamics at different locations on the protein surface of pol b. (Left) The scatter plots show the total solvation energy (Etot) and its three solvation components (E1–E3), all of which increases when the tryptophan probe is moving from interior to exterior. E1 completely disappears for buried mutants around 338 nm. The solid lines provide the guidelines for the trend. The gray bar indicates the dividing region around 338 nm separating mutants into buried and exposed groups. The inset sketch plot shows the spectral evolution with time. (Right) Three relaxation times correspond to different relaxation processes: hindered rotations of outer-layer bulk-type water in sub-picosecond (s1S); reorientation of inner-layer water in a few picoseconds (s2S); rearrangement of inner-layer hydration water rearrangement in tens-to-a hundred of picoseconds (s3S). The two slow relaxations times show strong correlations to the protein local properties such as charge density, hydrophobicity and steric confinement.
binding-site hydration water due to the enhanced confinement [44]. Therefore, hydration water around the pol b surface remains highly flexible on picosecond time scales in either apo or complex states. Retaining such high flexibility for both protein surface water and protein-DNA interfacial water is crucial for the polymerase function in mediating DNA binding and dissociation. 3.3. Femtosecond-resolved fluorescence anisotropy and protein sidechain dynamics To examine how the protein sidechain fluctuates during water relaxations, we also measured the tryptophan relaxations by its anisotropy dynamics. Fig. 5 shows the fluorescence transients at the parallel (I//) and perpendicular (I\) polarizations and the constructed fluorescence anisotropy r(t). Time-resolved fluorescence anisotropy gives the local molecular wobbling motions in picoseconds and the macromolecular tumbling motions in nanoseconds or longer. Here, with fs resolution we can resolve the fast wobbling motions of tryptophan in addition to the whole protein tumbling motion. As shown in Fig. 5, we observed four relaxation processes for either buried or exposed mutant. Taking the exposed mutant S109W as an example, the initial ultrafast relaxation happens in sIC <100 fs after deconvolution from the instrument response, due to the internal conversion (IC) from the 1Lb to the 1La state through conical intersection [45]. The slowest relaxation happens in sT 25 ns reflecting the whole enzyme tumbling motion. In between of the two extreme fast and slow processes exist two
more relaxations corresponding to the tryptophan wobbling motions from tens to hundreds of picoseconds (s2W = 21.7 ps and s3W = 156 ps). Similarly, we observed four relaxation processes for the buried mutant G179W with much slow wobbling dynamics (s2W = 34.1 ps and s3W = 381 ps). Generally, the first wobbling motion in tens of picoseconds must results from the indole ring reorientation while colliding with local hydration water molecules. The second wobbling motion in hundreds of picoseconds may result from the whole tryptophan wobbling motion, maybe including the backbone [46], coupled with the water-network reorganizations. In addition to the wobbling times, the anisotropy amplitude directly reports the wobbling space that can be further quantified by a semi-angle (h2W and h3W) in a wobbling cone model. All the wobbling semi-angles are small and in a range of 6.8–24° (see Table S2). Generally, the buried tryptophan takes a longer time to wobble in a highly constrained environment interacting with surrounding residues. The wobbling space is also much smaller than that of the exposed tryptophan. L22W, a special case of the exposed mutant, actually is buried and capped by the charged residues, significantly reducing its wobbling space (see supplementary material Table S2 and later discussion). 3.4. Correlations between hydration water dynamics and protein sidechain motions With a single tryptophan probe, we measured both the hydration dynamics and protein sidechain wobbling motions under the
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Fig. 5. Fluorescence anisotropy dynamics detected by tryptophan on pol b protein. (Top) Both parallel and perpendicular fluorescence transients were recorded. The inset shows the transients in a short time range. (Bottom) The constructed fluorescence anisotropy shows four decays including the initial ultrafast decay in sub-hundred femtosecond (sIC) due to the internal conversion, the two wobbling motions in tens (s2W) and hundreds of picoseconds (s3W), and the slow protein tumbling motion in nanoseconds (sT). The scatter plots show the experimental data and the solid lines are the best multi-exponential fits. The dashed lines mark each decay component.
same condition. This allows us to directly compare the protein sidechain dynamics to the local hydration-water relaxations as shown in Fig. 6A and B. Corresponding to the two solvation relaxations on picosecond time scales, the two wobbling motions happen on the same time scales but are much slower (siW > siS, i = 2, 3). This is true for all different sites on the protein surface, suggesting that hydration water is more flexible than the protein sidechains. Such flexible hydration water molecules may play a major role in controlling the protein dynamics as proposed by the ‘slaving’ model [13]. Recent studies on temperature dependence of water-protein coupled dynamics discovered that the protein sidechain motions are linearly correlated to the highly flexible hydration water dynamics [37,39]. These linear correlations can only be observed by altering temperature and hence cannot be observed in Fig. 6A and B due to the various water-protein coupling at different sites. Fortunately, the coupling strength reflected by the relaxation time ratio (ni = siW/siS, i = 2, 3) remains the same for all temperatures [37]. Fig. 6C plots the ratio at thirteen different sites on the protein surface and all the ratio values are larger than one. Surprisingly, the coupling ratio n2 is uniformly larger than n3, suggesting a stronger water-protein coupling on the hundreds of picoseconds time scale. Similar phenomenon was observed in the flexible polymerase Dpo4 [37]. Unlike water-protein rotational fluctuations around 20 ps, the water-protein coupling on hundreds of picoseconds can happen through both rotational and translational motions, which enhance water-protein coupling. From another point of view, the whole tryptophan reorientation requires large-scale higher cooperativity of water-network rearrangements. The ratio value ranges from 2 to 6 suggesting that
Fig. 6. Correlations between tryptophan wobbling and solvation times. (A and B) The tryptophan wobbling times are always longer than the corresponding solvation times. The dashed lines represent the equality of wobbling time and solvation time. (C) The ratio (ni = siW/s2S, i = 2, 3) signifies the coupling strength between hydration water and protein sidechains. It is always larger than one, reflecting the higher mobility of hydration water.
the protein sidechain only wobbles a few degrees before the completion of hydration water relaxation. Therefore, the protein movement is limited and the contributions in solvation energies are negligible, but the protein does participate in the relaxations.
3.5. Solvation speed, wobbling speed, hydration-water mobility and protein sidechain flexibility Due to distinct probe locations and protein surface heterogeneity, different mutants have different solvation energies and solvation times, and so do for the tryptophan wobbling motions. Therefore, we defined and calculated the solvation speed (Si = Ei/ siS, i = 1–3) and wobbling speed (xi = hi/siW, i = 2–3) and the results are shown in Fig. 7A and B, indicating the mobility of hydration water and flexibility of protein sidechains, respectively. The solvation speed S1 sharply increases with solvent exposure (kpeak) when the tryptophan probe moves towards the protein surface, detecting more outer-layer bulk-type hydration water. The solvation speed S2 also increases with solvent exposure but showing much less steep slope comparing to S1. This observation indicates that S2 includes partial contributions of the out-layer bulk-type water relaxing in a few picoseconds. This is in line with the bulk-water
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Fig. 7. (A) Solvation speed reports local hydration water mobility. (B) Tryptophan wobbling speed reports protein sidechain flexibility. Both solvation speed and tryptophan wobbling speed increase when the probe is moving from the protein interior to exterior. (C and D) The correlations between the wobbling speed and solvation speed at various locations suggest strong coupled water-protein interactions. The dashed circles mark the special position of L22W. As shown by the insets, L22W (yellow) is buried in the lyase domain where the local surface topology is highly confined. The surrounding charged residues further enhance the hydration water rigidity. The positive and negative charged residues are colored blue and red, respectively.
solvation occurring in 0.34 ps and 1.5 ps [37]. The solvation speed S3 quickly increases when the tryptophan probe is moving from the buried to exposed locations and then fluctuates for the exposed mutants. The highly wide distribution of S3 reflects the heterogeneous site-specific interactions between inner-layer hydration water and the protein. The wobbling speeds (x2 and x3) monotonically increase when the tryptophan probe is moving towards the surface possibly due to the coupled water-protein interactions because the increased water relaxation speed can drive the protein sidechain to fluctuate faster. Fig. 7C and D plot the correlations between the wobbling speeds and solvation speeds. The more labile the hydration water is, the more flexible the protein sidechain is. Our recent studies at various temperatures also confirmed the correlations between the wobbling speed and solvation speed, suggesting the direct water-protein coupling [37]. We also need to consider the steric confinement effects when the tryptophan probe is buried. As discussed before in 3.2 and 3.3, a unique case of L22W shows significantly slower relaxations for both the solvation and wobbling as shown in Fig. 7. The insets in Fig. 7C and D represent the local ribbon and surface structures, respectively. Firstly, 4 positive and 3 negative charged residues are nearby L22W, forming a strong electrostatic field on the local
hydration water. Secondly, the local surface structure shows a channel and deep clefts that easily trap water molecules. Thirdly, the L22W is located at the bottom of the channel and detects trapped hydration water. Therefore, the hydration dynamics detected by L22W is much slower. The slower wobbling dynamics of L22W accordingly results from the local structure inflexibility of both the protein and hydration water. L22W is in direct contact with hydrophobic residues on one side of the channel and on the other side of the channel interacts with four lysines residues that form a positively charged ring further stabilizing the indole ring of L22W. Finally, this unique charge distribution, constrained channel water and hydrophobic interactions together anchor the tryptophan probe in a limited space (see the smallest wobbling semi angles in Table S2 of the supplementary material). 4. Conclusions We have examined the protein surface hydration dynamics as well as related protein sidechain motions at 13 different locations of the apo state pol b enzyme by a tryptophan scan with femtosecond resolution. Two or three solvation components are identified for the protein with the buried or exposed tryptophan probe,
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respectively. When tryptophan is buried inside the protein, the probe only detects inner-layer hydration water that is typically within 7 Å from the protein surface. The solvation dynamics shows two relaxations occurring in a few picoseconds and tens-to-a hundred of picoseconds. For the exposed tryptophan on the protein surface, the probe detects both inner-layer and outer-layer water of the hydration shell, and the solvation dynamics shows an ultrafast sub-picosecond component and two relaxations on picosecond time scales. Generally, the picosecond solvation processes are much slower than that of bulk water and exhibit strong correlations with the protein surface properties such as hydrophobicity, charge density and local topology. We also observed two corresponding wobbling motions of the tryptophan sidechain on the similar time scales but they are always slower than hydrationwater relaxations. The protein sidechain wobbling space is generally small and even smaller for the buried environment. Therefore, the protein displacement seems negligible when the water relaxation happens. The solvation energy distribution further suggests that solvation is mainly due to the hydration-water response and the protein contribution is negligible. The sub-picosecond solvation is completely due to the relaxation of out-layer bulk-type hydration water. The two slow water relaxations from a few to a hundred picoseconds result from the collective water reorientation and water-network rearrangement. The highly constrained inner-layer hydration water makes dominant contributions to the two slow relaxations while the outer-layer hydration water contributes partially to the solvation in a few picoseconds. Consistent with the strong waterprotein interactions, the two wobbling dynamics seems driven by the interfacial, inner-layer hydration water relaxations.
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Acknowledgements It was the experience of working in the Zewail group about 17 years ago that first triggered my interest in hydration water and protein dynamics. Further collaboration with Dr. Zewail in this area, and his brilliant vision, insights, and clarity on the complexity of hydration dynamics have reshaped our thinking of the protein hydration field. We thank Prof. Ming-Daw Tsai’s group for kindly providing the plasmid of rat pol b (PET17b) and also thank Dr. Jin Yang for discussion. This work was supported in part by the National Institute of Health Grants GM095997 and GM118332. The MD simulations were supported in part by an allocation of computing time through the Ohio Supercomputer Center. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.cplett.2017.03. 002. References [1] F. Franks, Royal Society of Chemistry (Great Britain). Water: A matrix of life, Royal Society of Chemistry, Royal Society of Chemistry, Cambridge, 2000. [2] R. Zhou, X. Huang, C.J. Margulis, B.J. Berne, Hydrophobic collapse in multidomain protein folding, Science 305 (2004) 1605–1609. [3] Y. Levy, J.N. Onuchic, Water mediation in protein folding and molecular recognition, Annu. Rev. Biophys. Biomol. Struct. 35 (2006) 389–415. [4] S.J. Kim, B. Born, M. Havenith, M. Gruebele, Real-time detection of proteinwater dynamics upon protein folding by terahertz absorption spectroscopy, Angew. Chem. Int. Ed. 47 (2008) 6486–6489. [5] D. Zhong, A. Douhal, A.H. Zewail, Femtosecond studies of protein-ligand hydrophobic binding and dynamics: human serum albumin, Proc. Natl. Acad. Sci. USA 97 (2000) 14056–14061. [6] B. Jayaram, T. Jain, The role of water in protein-DNA recognition, Annu. Rev. Biophys. Biomol. Struct. 33 (2004) 343–361. [7] A. Ghosh, J. Wang, Y.S. Moroz, I.V. Korendovych, M. Zanni, W.F. DeGrado, F. Gai, R.M. Hochstrasser, 2D IR spectroscopy reveals the role of water in the binding
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Research paper
Dynamics of hydration water and coupled protein sidechains around a polymerase protein surface Yangzhong Qin, Yi Yang, Lijuan Wang, Dongping Zhong ⇑ Department of Physics, Department of Chemistry and Biochemistry, Programs of Biophysics, Chemical Physics, and Biochemistry, The Ohio State University, Columbus, OH 43210, United States
a r t i c l e
i n f o
Article history: Received 31 December 2016 In final form 2 March 2017 Available online xxxx Keywords: Hydration dynamics Slaved sidechain motions Water-protein coupled fluctuations Femtosecond spectroscopy Tryptophan scan
a b s t r a c t Water-protein coupled interactions are essential to the protein structural stability, flexibility and dynamic functions. The ultimate effects of the hydration dynamics on the protein fluctuations remain substantially unexplored. Here, we investigated the dynamics of both hydration water and protein sidechains at 13 different sites around the polymerase b protein surface using a tryptophan scan with femtosecond spectroscopy. Three types of hydration-water relaxations and two types of protein sidechain motions were determined, reflecting a highly dynamic water-protein interactions fluctuating on the picosecond time scales. The hydration-water dynamics dominate the coupled interactions with higher flexibility. Ó 2017 Elsevier B.V. All rights reserved.
1. Introduction Water is a necessity for nearly all life known [1]. It involves in most biological processes not only as a universal solvent, but also as an active participant in mediating protein folding [2–4], protein-ligand binding [5–7] and enzymatic catalysis [8–10]. Hydration water, directly interacting with protein at the waterprotein interface, strongly influences the protein structure and dynamics [11–15]. Mutually, the water structure and dynamics are significantly perturbed as well [16–21]. Understanding the dynamics of coupled water-protein interactions is crucial to revealing its ultimate effects on the protein’s functions that require both local and global flexibility in aqueous solution. Extensive studies had been carried out to investigate the dynamics of either hydration water or proteins by both experiments [22–28] and simulation [14,20,21,29]. However, due to the strong water-protein coupling, it remains challenging to disentangle their dynamics. Here, we report our simultaneous measurements of the surface hydration dynamics and related protein sidechain motions by employing the femtosecond (fs)-resolved fluorescence spectroscopy and a tryptophan (W) scan [18,19]. With site-directed mutagenesis, we placed a single tryptophan, one at a time, as an optical probe at 13 different locations on the surface of DNA polymerase b (Pol b) to globally map out the dynamics of hydration water and protein sidechains (Fig. 1). The structure and function ⇑ Corresponding author. E-mail address: [email protected] (D. Zhong).
of Pol b have been extensively studied previously [30–34]. As an X-family DNA polymerase, it can bind to the gapped or nicked DNA and fulfill gap filling synthesis. Pol b has 335 amino acids (39 kDa) with the overall structure following the right hand architecture (Fig. 1). The polymerase domain (31 kDa) consists of three subdomains: a double-stranded DNA (dsDNA) binding subdomain D (also called thumb), a nucleotidyl-transfer subdomain C (palm), and a dNTP selection subdomain N (finger). The N-terminal lyase domain (8 kDa) performs the lyase function to remove the 50 -deoxyribose phosphate during base excision repair [32]. All tryptophan probes were scattered on different domains of Pol b to maximally sample the protein surface heterogeneity. Meanwhile, all tryptophan probes except W325 (WT) are designed at or close to the DNA binding sites that can examine the hydration dynamics of binding sites contrasting to non-binding sites in the apo state [31]. 2. Materials and methods The plasmid of rat pol b (PET17b) was kindly provided by the Tsai’s group at the Ohio State University. Wild type pol b contains a single tryptophan at W325, which was mutated to W325F and served as the double mutant template. Twelve double mutants, each with a single tryptophan, were designed. The plasmids of wild type and all twelve double mutants were overexpressed in Escherichia coli BL21(DE3) one by one. Each protein was then purified following previously reported protocol [34]. We further checked the CD spectra and confirmed the similar structures of the mutants
http://dx.doi.org/10.1016/j.cplett.2017.03.002 0009-2614/Ó 2017 Elsevier B.V. All rights reserved.
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Fig. 1. Ribbon and surface structures of pol b in the apo state (Protein Data Bank ID: 1BPD). Pol b includes a lyase domain (gray) and three polymerase subdomains, thumb (green), palm (red), and finger (blue), forming a right-hand configuration. Yellow spheres and patches indicate the positions of tryptophan probes. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
to that of WT. Each protein was finally stored in a buffer containing 100 mM Tris-HCl (pH 8.0), 150 mM KCl, 2 mM DTT, 3% glycerol (vol/vol), and 1.5 mM MgCl2. The final protein concentration is about 400 lM. The complete details of the time-resolved fluorescence spectroscopy were reported previously [35]. In summary, we used the fluorescence up-conversion technique. An 800 nm pulsed laser beam with 1 kHz repletion rate and 110 fs temporal width was generated by a commercial amplifier (Spitfire, Spectra-Physics). It was split into two parts, with 10% to serve as the probe beam and 90% to pump an optical parametric amplifier (OPA 800C, Spectra-Physics). The OPA generated 590-nm laser pulses that traveled through the translation stage with maximum 3 ns delay relative to the probe. Then, it was focused on a doubling BBO crystal to produce 295 nm pump pulses. Each pump pulse energy was attenuated to 140 nJ before focusing on the protein sample stored in a motor-controlled spinning cell with 1 mm thickness. The protein fluorescence was collected and focused on the up-conversion BBO crystal to mix with a focused probe pulse. The up-converted signal was directed to a monochromator coupled with a photomultiplier tube (PMT). The instrument response time under current non-collinear condition is around 450 fs determined by water Raman signal at 325 nm. For all the solvation transients, the pump pulse polarization was set to the magic angle (54.7°) with respect to the acceptance axis of the up-conversion BBO crystal. For fluorescence anisotropy study, the polarization of pump pulse was set to be either parallel or perpendicular to the acceptance axis to record the parallel signal (I//) and perpendicular signal (I\), respectively. The probe beam polarization was always parallel to the acceptance axis of the up-conversion BBO crystal. 3. Results and discussion 3.1. Femtosecond-resolved fluorescence transient dynamics For each mutant, we have measured the fluorescence transients with a 3 ns time window at different wavelengths from 310 nm to 370 nm covering the blue- and red-side emission spectrum. Fig. 2 shows the typical fluorescence transients of a buried mutant G179W and an exposed mutant S109W. Detailed data-analysis strategy was provided previously [35]. We observed typical solvation patterns for all mutants. For the buried mutant G179W, beside two lifetime decays in nanoseconds (1.2 ns and 5.5 ns), we observed two picosecond solvation decays (4.5–11.2 ps and 82– 108 ps) at the blue side and one rise component (3.8–7.0 ps) at the red side of the emission spectrum. For the exposed mutant S109W, we observed three solvation decays (0.38–0.6 ps, 3.6– 5.0 ps and 34–51 ps) at the blue side and one rise component
Fig. 2. Normalized fluorescence transients at a series of wavelengths for the buried mutant G179W (A) and exposed mutant S109W (B). The scatter plots and the solid lines show the raw data and best multi-exponential fits, respectively.
(0.8–1.1 ps) at the red side in addition to the two lifetime decays (0.7 ns and 5.1 ns). All these decay and rise dynamics are typical solvation signature, which has been observed in many other proteins [18,19], reflecting the relaxations of excited tryptophan and its surroundings. 3.2. Solvent exposure, solvation correlation function and hydration dynamics To further understand the solvation dynamics measured by tryptophan at different locations, we need to know the environ-
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ment of tryptophan probe such as its local protein topology and solvent exposure. Fig. 3A presents a snapshot of MD simulations showing the protein structure as well as hydration water detected by tryptophan for mutant G179W and S109W. The steady-state emission peak can roughly serve as an indicator for the solvent exposure. Fig. 3B shows the steady-state emission spectra of selected four mutants. Bluer emission indicates buried environment of tryptophan and less solvent exposure while redder emission indicates the exposed position of tryptophan and higher solvent exposure [36]. The gradual change of solvent exposure can be approximately quantified by the steady-state emission peaks listed in Table S1 of the supplementary material. Typically, with the dividing line around 338 nm of the emission peak (kpeak) [36], we can separate different mutants into two groups: G179W and F146W as a buried group and all the others as an exposed group. With the fluorescence transients gated at different wavelengths, one can construct the fs-resolved emission spectra (FRES) at different times and monitor the dynamic Stokes shifts with time. As demonstrated in previous studies [18,19], the solvation correlation
Fig. 3. (A) A snapshot of MD simulations shows the water molecules detected within 10 Å from the indole ring of tryptophan for the buried mutant G179W and exposed mutant S109W. Buried tryptophan only detects inner-layer (red) hydration water within 7 Å from the protein surface and exposed tryptophan detects both inner- and outer-layer (blue) hydration water. (B) The steady-state emission spectrum gradually shifts to the red side when the tryptophan is moving from a buried to exposed position, with a dividing line around 338 nm. (C) The solvation correlation functions report either two or three water relaxation processes by the buried mutant G179W and exposed mutant S109W, respectively.
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function C(t) can be deduced, which directly measures the solvation relaxation. Fig. 3C shows the solvation correlation functions for both the buried mutant G179W and exposed mutant S109W. Consistent with fluorescence transients, the solvation correlation function exhibits only two decays (6.7 ps and 106 ps) for the buried mutant and three decays (0.53 ps, 3.9 ps and 47 ps) for the exposed mutant (see Table S1 for details). The ultrafast component detected by the exposed mutant comes from the outer-layer water of the hydration shell that is distant from the protein surface. Indeed, our recent studies in many proteins suggest that buried mutants only detect hydration water within 7 Å of the inner hydration layers from the protein surface [37–39], but exposed mutants can detect water molecules beyond 7 Å forming the outer hydration layers as shown in Fig. 3A. Fig. 4 shows the distributions of solvation energies and solvation relaxation times of all mutants. The total solvation energy (Etot) and the ultrafast solvation energy (E1) gradually increase when the solvent exposure is increasing. The E2 and E3 increase before the emission peaks reach 338 nm and stay nearly no change afterwards. This suggests that both E2 and E3, especially E3, mainly come from the inner-layer hydration water response. It also suggests that the protein solvation contribution may be negligible, consistent with our recent studies [37–39], because both E2 and E3 keep decreasing when solvent exposure reduces. Detailed solvation energies and solvation times are listed in Table S1. The ultrafast solvation happens in sub-picosecond for all exposed mutants, close to the bulk-water relaxation around 0.34 ps [37]. Therefore, it reflects the bulk-type nature of the outer-layer water of the hydration shell. On such an ultrafast time scale, the solvation mainly comes from the water libration and hindered rotation. The second solvation occurs in 3.9–8.7 ps (s2S) during which water molecules typically undergo large amplitude reorientations [40,41]. The slowest relaxation happens in 47–106 picoseconds, reflecting the whole water network reorganizations, coupled with local protein fluctuations. This time scale is consistent with the surface hydration diffusion time and also residence time reported by NMR studies as well as by simulations [24,42,43]. Due to the close interactions of inner-layer hydration water with the protein, the slow relaxation times show clear distinctions that are correlated to the protein surface heterogeneity. For an example, hydration water around hydrophobic sites (S109W and K41W) show relatively faster relaxation times for both water reorientations and water network reorganizations. The heavily charged environment shows the slowest relaxations. Local protein steric confinement also plays an important role in slowing down the hydration-water relaxation. Buried mutants of F146W and G179W basically detect inner-layer hydration water molecules that are confined on the rugged protein surface. Although L22W is categorized as an exposed mutant, the local protein surface forms deep pockets and clefts, rendering high restriction to the inner-layer hydration water and hence causing water molecules to move slower. All these results lead us to the conclusion that the protein surface hydration dynamics are directly coupled to the protein local environments such as hydrophobicity, charge density and steric confinement. This is in agreement with previous studies both experimentally [35] and computationally [41]. Except WT, all other mutants were designed to have the tryptophan probe placed around the DNA binding site. The WT protein serves as a control to report the hydration dynamics at nonbinding sites and all other mutants report the hydration dynamics at the DNA binding sites. As shown in Fig. 4, no clear distinctions were observed for the hydration dynamics at the DNA binding sites compared to that of the non-binding site (WT). However, this is only true for the apo state without binding to DNA. Previous studies on the binary (pol b-DNA) and ternary (pol b-DNA-dNTP) complex states showed a moderate slowdown by a factor of 2 for the
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Fig. 4. Solvation dynamics at different locations on the protein surface of pol b. (Left) The scatter plots show the total solvation energy (Etot) and its three solvation components (E1–E3), all of which increases when the tryptophan probe is moving from interior to exterior. E1 completely disappears for buried mutants around 338 nm. The solid lines provide the guidelines for the trend. The gray bar indicates the dividing region around 338 nm separating mutants into buried and exposed groups. The inset sketch plot shows the spectral evolution with time. (Right) Three relaxation times correspond to different relaxation processes: hindered rotations of outer-layer bulk-type water in sub-picosecond (s1S); reorientation of inner-layer water in a few picoseconds (s2S); rearrangement of inner-layer hydration water rearrangement in tens-to-a hundred of picoseconds (s3S). The two slow relaxations times show strong correlations to the protein local properties such as charge density, hydrophobicity and steric confinement.
binding-site hydration water due to the enhanced confinement [44]. Therefore, hydration water around the pol b surface remains highly flexible on picosecond time scales in either apo or complex states. Retaining such high flexibility for both protein surface water and protein-DNA interfacial water is crucial for the polymerase function in mediating DNA binding and dissociation. 3.3. Femtosecond-resolved fluorescence anisotropy and protein sidechain dynamics To examine how the protein sidechain fluctuates during water relaxations, we also measured the tryptophan relaxations by its anisotropy dynamics. Fig. 5 shows the fluorescence transients at the parallel (I//) and perpendicular (I\) polarizations and the constructed fluorescence anisotropy r(t). Time-resolved fluorescence anisotropy gives the local molecular wobbling motions in picoseconds and the macromolecular tumbling motions in nanoseconds or longer. Here, with fs resolution we can resolve the fast wobbling motions of tryptophan in addition to the whole protein tumbling motion. As shown in Fig. 5, we observed four relaxation processes for either buried or exposed mutant. Taking the exposed mutant S109W as an example, the initial ultrafast relaxation happens in sIC <100 fs after deconvolution from the instrument response, due to the internal conversion (IC) from the 1Lb to the 1La state through conical intersection [45]. The slowest relaxation happens in sT 25 ns reflecting the whole enzyme tumbling motion. In between of the two extreme fast and slow processes exist two
more relaxations corresponding to the tryptophan wobbling motions from tens to hundreds of picoseconds (s2W = 21.7 ps and s3W = 156 ps). Similarly, we observed four relaxation processes for the buried mutant G179W with much slow wobbling dynamics (s2W = 34.1 ps and s3W = 381 ps). Generally, the first wobbling motion in tens of picoseconds must results from the indole ring reorientation while colliding with local hydration water molecules. The second wobbling motion in hundreds of picoseconds may result from the whole tryptophan wobbling motion, maybe including the backbone [46], coupled with the water-network reorganizations. In addition to the wobbling times, the anisotropy amplitude directly reports the wobbling space that can be further quantified by a semi-angle (h2W and h3W) in a wobbling cone model. All the wobbling semi-angles are small and in a range of 6.8–24° (see Table S2). Generally, the buried tryptophan takes a longer time to wobble in a highly constrained environment interacting with surrounding residues. The wobbling space is also much smaller than that of the exposed tryptophan. L22W, a special case of the exposed mutant, actually is buried and capped by the charged residues, significantly reducing its wobbling space (see supplementary material Table S2 and later discussion). 3.4. Correlations between hydration water dynamics and protein sidechain motions With a single tryptophan probe, we measured both the hydration dynamics and protein sidechain wobbling motions under the
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Fig. 5. Fluorescence anisotropy dynamics detected by tryptophan on pol b protein. (Top) Both parallel and perpendicular fluorescence transients were recorded. The inset shows the transients in a short time range. (Bottom) The constructed fluorescence anisotropy shows four decays including the initial ultrafast decay in sub-hundred femtosecond (sIC) due to the internal conversion, the two wobbling motions in tens (s2W) and hundreds of picoseconds (s3W), and the slow protein tumbling motion in nanoseconds (sT). The scatter plots show the experimental data and the solid lines are the best multi-exponential fits. The dashed lines mark each decay component.
same condition. This allows us to directly compare the protein sidechain dynamics to the local hydration-water relaxations as shown in Fig. 6A and B. Corresponding to the two solvation relaxations on picosecond time scales, the two wobbling motions happen on the same time scales but are much slower (siW > siS, i = 2, 3). This is true for all different sites on the protein surface, suggesting that hydration water is more flexible than the protein sidechains. Such flexible hydration water molecules may play a major role in controlling the protein dynamics as proposed by the ‘slaving’ model [13]. Recent studies on temperature dependence of water-protein coupled dynamics discovered that the protein sidechain motions are linearly correlated to the highly flexible hydration water dynamics [37,39]. These linear correlations can only be observed by altering temperature and hence cannot be observed in Fig. 6A and B due to the various water-protein coupling at different sites. Fortunately, the coupling strength reflected by the relaxation time ratio (ni = siW/siS, i = 2, 3) remains the same for all temperatures [37]. Fig. 6C plots the ratio at thirteen different sites on the protein surface and all the ratio values are larger than one. Surprisingly, the coupling ratio n2 is uniformly larger than n3, suggesting a stronger water-protein coupling on the hundreds of picoseconds time scale. Similar phenomenon was observed in the flexible polymerase Dpo4 [37]. Unlike water-protein rotational fluctuations around 20 ps, the water-protein coupling on hundreds of picoseconds can happen through both rotational and translational motions, which enhance water-protein coupling. From another point of view, the whole tryptophan reorientation requires large-scale higher cooperativity of water-network rearrangements. The ratio value ranges from 2 to 6 suggesting that
Fig. 6. Correlations between tryptophan wobbling and solvation times. (A and B) The tryptophan wobbling times are always longer than the corresponding solvation times. The dashed lines represent the equality of wobbling time and solvation time. (C) The ratio (ni = siW/s2S, i = 2, 3) signifies the coupling strength between hydration water and protein sidechains. It is always larger than one, reflecting the higher mobility of hydration water.
the protein sidechain only wobbles a few degrees before the completion of hydration water relaxation. Therefore, the protein movement is limited and the contributions in solvation energies are negligible, but the protein does participate in the relaxations.
3.5. Solvation speed, wobbling speed, hydration-water mobility and protein sidechain flexibility Due to distinct probe locations and protein surface heterogeneity, different mutants have different solvation energies and solvation times, and so do for the tryptophan wobbling motions. Therefore, we defined and calculated the solvation speed (Si = Ei/ siS, i = 1–3) and wobbling speed (xi = hi/siW, i = 2–3) and the results are shown in Fig. 7A and B, indicating the mobility of hydration water and flexibility of protein sidechains, respectively. The solvation speed S1 sharply increases with solvent exposure (kpeak) when the tryptophan probe moves towards the protein surface, detecting more outer-layer bulk-type hydration water. The solvation speed S2 also increases with solvent exposure but showing much less steep slope comparing to S1. This observation indicates that S2 includes partial contributions of the out-layer bulk-type water relaxing in a few picoseconds. This is in line with the bulk-water
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Fig. 7. (A) Solvation speed reports local hydration water mobility. (B) Tryptophan wobbling speed reports protein sidechain flexibility. Both solvation speed and tryptophan wobbling speed increase when the probe is moving from the protein interior to exterior. (C and D) The correlations between the wobbling speed and solvation speed at various locations suggest strong coupled water-protein interactions. The dashed circles mark the special position of L22W. As shown by the insets, L22W (yellow) is buried in the lyase domain where the local surface topology is highly confined. The surrounding charged residues further enhance the hydration water rigidity. The positive and negative charged residues are colored blue and red, respectively.
solvation occurring in 0.34 ps and 1.5 ps [37]. The solvation speed S3 quickly increases when the tryptophan probe is moving from the buried to exposed locations and then fluctuates for the exposed mutants. The highly wide distribution of S3 reflects the heterogeneous site-specific interactions between inner-layer hydration water and the protein. The wobbling speeds (x2 and x3) monotonically increase when the tryptophan probe is moving towards the surface possibly due to the coupled water-protein interactions because the increased water relaxation speed can drive the protein sidechain to fluctuate faster. Fig. 7C and D plot the correlations between the wobbling speeds and solvation speeds. The more labile the hydration water is, the more flexible the protein sidechain is. Our recent studies at various temperatures also confirmed the correlations between the wobbling speed and solvation speed, suggesting the direct water-protein coupling [37]. We also need to consider the steric confinement effects when the tryptophan probe is buried. As discussed before in 3.2 and 3.3, a unique case of L22W shows significantly slower relaxations for both the solvation and wobbling as shown in Fig. 7. The insets in Fig. 7C and D represent the local ribbon and surface structures, respectively. Firstly, 4 positive and 3 negative charged residues are nearby L22W, forming a strong electrostatic field on the local
hydration water. Secondly, the local surface structure shows a channel and deep clefts that easily trap water molecules. Thirdly, the L22W is located at the bottom of the channel and detects trapped hydration water. Therefore, the hydration dynamics detected by L22W is much slower. The slower wobbling dynamics of L22W accordingly results from the local structure inflexibility of both the protein and hydration water. L22W is in direct contact with hydrophobic residues on one side of the channel and on the other side of the channel interacts with four lysines residues that form a positively charged ring further stabilizing the indole ring of L22W. Finally, this unique charge distribution, constrained channel water and hydrophobic interactions together anchor the tryptophan probe in a limited space (see the smallest wobbling semi angles in Table S2 of the supplementary material). 4. Conclusions We have examined the protein surface hydration dynamics as well as related protein sidechain motions at 13 different locations of the apo state pol b enzyme by a tryptophan scan with femtosecond resolution. Two or three solvation components are identified for the protein with the buried or exposed tryptophan probe,
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respectively. When tryptophan is buried inside the protein, the probe only detects inner-layer hydration water that is typically within 7 Å from the protein surface. The solvation dynamics shows two relaxations occurring in a few picoseconds and tens-to-a hundred of picoseconds. For the exposed tryptophan on the protein surface, the probe detects both inner-layer and outer-layer water of the hydration shell, and the solvation dynamics shows an ultrafast sub-picosecond component and two relaxations on picosecond time scales. Generally, the picosecond solvation processes are much slower than that of bulk water and exhibit strong correlations with the protein surface properties such as hydrophobicity, charge density and local topology. We also observed two corresponding wobbling motions of the tryptophan sidechain on the similar time scales but they are always slower than hydrationwater relaxations. The protein sidechain wobbling space is generally small and even smaller for the buried environment. Therefore, the protein displacement seems negligible when the water relaxation happens. The solvation energy distribution further suggests that solvation is mainly due to the hydration-water response and the protein contribution is negligible. The sub-picosecond solvation is completely due to the relaxation of out-layer bulk-type hydration water. The two slow water relaxations from a few to a hundred picoseconds result from the collective water reorientation and water-network rearrangement. The highly constrained inner-layer hydration water makes dominant contributions to the two slow relaxations while the outer-layer hydration water contributes partially to the solvation in a few picoseconds. Consistent with the strong waterprotein interactions, the two wobbling dynamics seems driven by the interfacial, inner-layer hydration water relaxations.
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