Dynamic Solvation and Coupling of the Hydration Shell of ZnII

Article pubs.acs.org/JPCB

Dynamic Solvation and Coupling of the Hydration Shell of ZnII-Substituted Cytochrome c in the Presence of Guanidinium Ions Jagnyaseni Tripathy,† Jenny Jo Mueller,‡ Nolan C. Shepherd,§ and Warren F. Beck* Department of Chemistry, Michigan State University, East Lansing, Michigan 48824, United States ABSTRACT: The fluorescence Stokes shift (FSS) response of ZnII-substituted cytochrome c (ZnCytc) is transformed from a monotonic red-shifting response in water to a bidirectional response with much slower time constants in the presence of low concentrations of guanidinium (Gdm+) ions. The FSS response in water observed over the 100 ps to 10 ns range has two exponential components with time constants of 135 ps and 1.6 ns that account for a total shift of 30 cm−1, about onehalf of the solvation reorganization energy. In contrast, in the presence of only 0.25 M Gdm+, the FSS response initially shifts 21 cm−1 to the blue with a 820 ps time constant and then shifts 60 cm−1 back to the red with a 3.5 ns time constant. The effect of Gdm+ on the FSS response effectively saturates at 1.0 M, well below the 1.75 M midpoint of the two-state unfolding transition. These results establish that the FSS response in ZnCytc includes a significant contribution from the surrounding hydration shell, which assumes a perturbed hydrogen-bonding network owing to the binding of Gdm+ ions to the protein surface. The blue-shifting part of the FSS response arises from a light-induced conformational change that expands the protein- and solvent-derived cavity around the excited-state ZnII porphyrin. This non-polar part of the solvation response is enhanced in the presence of Gdm+ because the protein/solvent surroundings of the ZnII porphyrin are effectively more flexible than in water. The enhanced flexibility in the presence of Gdm+ increases the amplitudes and accordingly lengthens the correlation time scales for the protein and hydration-shell fluctuations that contribute to the FSS response.

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INTRODUCTION The hydration shell of a protein constitutes a domain of “biological water”1−4 that contributes to the stability of the native folded structure5−8 and to its biological function, especially in charge transfer or redox catalysis,9 owing to its distinct dynamical properties compared to the bulk.10−13 The thickness of the hydration shell inferred by Gruebele and coworkers using terahertz spectroscopy is at least 1.0 nm.14 In work by Zewail and co-workers,2,3,15 femtosecond timeresolved fluorescence measurements using probe chromophores on or tethered close to a protein’s surface showed that the diffusive part of the polar solvation response is slowed at least to the 10−100 ps time scale from the sub-ps regime that is characteristic of bulk water.16 A comparable slowing of reorientational dynamics occurs in water near interfaces,17,18 but the solvation response detected by intrinsic or proteinbound probes reaches well into the nanosecond regime.19−26 Such long time scales are characteristic of protein-derived motions and depend on the protein-folding state,27 but the contribution of the hydration layer to the detected response is not easily distinguished from that of the protein.9,28−30 In fact, molecular dynamics simulations by Matyushov and co-workers show that slow motions of water molecules in the hydration shell on the ns time scale arise from electrostatic interactions with surface charges and mechanical coupling (or “slaving” 31−33) to conformational motions of the solvated protein.9,34−37 It follows that the hydration shell should also be expected to contribute strongly to the rate at which a protein © 2013 American Chemical Society

evolves structurally as it descends toward the native structure from the unfolded or denatured state.38 In this paper, the coupling of the hydration shell with the underlying protein structure is considered in a study of the changes in protein dynamics that occur as ZnII-substituted cytochrome c (ZnCytc)39,40 is perturbed by the addition of guanidinium (Gdm+) ions. The ZnII porphyrin in ZnCytc (see Figure 1) is used as an intrinsic electronic probe that reports protein-derived motions over the 100 ps to 20 ns regime via its fluorescence Stokes shift (FSS) response function. The plan was to determine how the FSS response from ZnCytc changes as the addition of Gdm+ shifts the equilibrium ensemble toward the unfolded state. The action of Gdm+ on the stability of a folded protein is associated with binding to surface peptide linkages41 or to changes in the hydrogen-bonding structure of water near the protein−water interface or on its surface.42 Thus, if the motions of the protein are coupled to those of the hydration shell, the FSS response sensed by the ZnII porphyrin should be transformed by the addition of Gdm+. The results presented here confirm this expectation, but the surprise is that an unusual bidirectional FSS response is observed even at very low Gdm+ concentrations, well below the unfolding transition, and the response is slowed by about a factor of 2 compared to that in water. Received: May 7, 2013 Revised: September 16, 2013 Published: October 22, 2013 14589

dx.doi.org/10.1021/jp404554t | J. Phys. Chem. B 2013, 117, 14589−14598

The Journal of Physical Chemistry B

Article

that exhibits large-amplitude fluctuations is formed upon demetalation of the porphyrin,58 the two exponential components in the FSS response from metal-free cytochrome c (fbCytc) are nearly 10 times longer than those of ZnCytc.27 This comparison of the FSS response from ZnCytc with that of fbCytc strongly suggests that protein-derived motions dominantly contribute to the two correlation times observed on the fluorescence time scale.27 The new results we present here indicate that the hydration shell also contributes strongly to the FSS response of ZnCytc. Even at low Gdm+ concentrations and in the absence of a significant fraction of unfolded protein in the ensemble, an unusual bidirectional and slowed FSS response is observed. The results indicate that binding of Gdm+ to the protein surface transforms the FSS response by altering the hydrogen-bonding network of the surrounding hydration shell and enhancing the flexibility of the coupled protein−solvent interface.

Figure 1. Ribbon (left) and solvent-excluded surface (right) renderings of the X-ray crystal structure of horse-heart ferricytochrome c (1hrc.pdb).43 The porphyrin, methionine (M80), and histidine (H18) residues that serve as axial ligands to the FeIII ion and the cysteine residues (C14 and C17) that link the porphyrin to the polypeptide backbone are shown as stick structures in the ribbon picture. The protein structure is color coded from red to blue in order of relative folding stability following the scheme of Englander and coworkers:44 residues 70−85 (red), residues 36−61 (yellow), residues 20−35 and the α helix over residues 60−70 (green), and the N- and C-terminal α helices (blue). On the basis of a comparison of 2D NMR spectra, ZnCytc is isostructural with the native, FeII-containing protein in solution.45,46

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EXPERIMENTAL SECTION Sample Preparation. ZnCytc was prepared from horseheart ferricytochrome c (Sigma) after demetalation in liquid anhydrous HF using the approach described by Vanderkooi and co-workers39,40 and the chromatographic procedures described by Winkler and co-workers59 and Kostić and coworkers.60 The metal-free intermediate (fbCytc) was reconstituted at 50 °C with ZnII in the presence of a 10-fold molar excess of zinc acetate. Formation of the ZnII porphyrin was followed spectrophotometrically by monitoring the Q-band vibronic structure, which evolves from the four-peak structure characteristic of the metal-free porphyrin to the two-peak structure of the metalloporphyrin. The ZnCytc product was concentrated using several ultrafiltration and redilution cycles in a 25 mM phosphate buffer solution at pH 7.0 over an Amicon (Millipore) YM10 membrane. The concentrated product solution was flash frozen in liquid nitrogen and stored in a −85 °C freezer. For use in fluorescence experiments, frozen samples of ZnCytc were thawed and then diluted with a 25 mM phosphate buffer solution at pH 7.0 containing a selected GdmCl concentration (0.25−5 M). After correction of the pH to 7.0 and incubation at room temperature in darkness for 1 h, the solution was passed through a 0.22 μm microfilter. The final concentration of the sample was then adjusted with pH 7.0 GdmCl solution to obtain an absorption of 0.1−0.2 at 584 nm, the maximum of the Q-band 0−0 peak, for a path length of 1.0 cm. The sample was held in a quartz cuvette (1 cm path), and the headspace above the sample was purged with nitrogen gas just prior to an experiment. In the picosecond fluorescence spectrometer, the sample cuvette was held in a water-cooled holder that was maintained at 22 °C using a Neslab RTE-110 water bath. Continuous-Wave Absorption and Fluorescence Spectroscopy. Absorption spectra were acquired with a Hitachi U-2000 spectrophotometer (2 nm band-pass). Fluorescence spectra were obtained with a home-built spectrofluorimeter58 consisting of a Jobin-Yvon AH10 100 W tungsten-halogen light source, a Jobin-Yvon H10 excitation monochromator (4 nm bandpass), an Acton Research SP-150 emission spectrograph (2 nm bandpass), and a Jobin-Yvon Symphony charge-coupled device (CCD) detector. The CCD detector employs a liquid nitrogen cooled, back-illuminated, 2000 × 800 pixel silicon detector chip (EEV corporation). The sample cuvette was held in a Quantum Northwest TLC50F

In previous work in our laboratory,23,47 we characterized the FSS response from ZnCytc in the native state in water and in mixtures of glycerol and water. The FSS response function, Sv(t), is experimentally determined as Sv(t ) =

v (t ) − v (∞ ) v(0) − v(∞)

(1)

where v(t) relates the mean frequency of the time-resolved fluorescence spectrum at a given instant in time following optical excitation of the probe. In the linear-response regime, where the fluctuation−dissipation relation is applicable, the FSS response is equivalent to the time-correlation function for the probe’s optical transition, M(t), M (t ) =

⟨Δω(0)Δω(t )⟩ ⟨(Δω)2 ⟩

(2)

which is determined from the time-averaged and instantaneous ground-to-excited-state transition frequencies, ⟨ω⟩ and ω(t), respectively, and the fluctuation, Δω(t) = ⟨ω⟩ − ω(t), with averaging over the ensemble.48,49 The polar part of the FSS response arises from the dielectric relaxation of the surrounding medium in response to the probe chromophore’s excited-state change in permanent dipole moment.50 On the ps time scale, the polar solvation response in molecular solvents and in chromoproteins corresponds to rotational diffusion;51 on the sub-ps time scale, hindered rotations (librations) and inertial (free-rotor) motions make large contributions to the response.52−55 The non-polar part of the FSS response arises from the viscoelastic response of the solvent to the changes in the probe’s size that accompany the optical transition.56,57 The FSS response from ZnCytc in water consists of a monotonic red shift of the time-resolved fluorescence spectrum with two exponential components. The time constants, 250 ps and 1.5 ns, were assigned to fluctuations of the hydrophobic core and of solvent-contacting regions of the protein, respectively, owing to the weak glycerol dependence of the latter time constant.23 Because a molten-globule-like structure 14590



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preparations used in this study, which were worked up using ion-exchange chromatography,59,60 reproducibly exhibit significantly narrower absorption and fluorescence line shapes than observed previously23 in preparations isolated by gel-filtration chromatography.39,40 The solvation reorganization energy, λ,54 as estimated from the frequency shift between the absorption (A) and fluorescence (F) 0−0 peaks,

Peltier-effect temperature controller; the temperature set point was 22 °C. The absorption and fluorescence instruments were controlled by LabVIEW (National Instruments) programs. Picosecond Fluorescence Spectroscopy. Time-resolved fluorescence spectra were acquired using a two-channel, Tformat, time-correlated single photon counting (TCSPC) instrument. Excitation pulses (tuned over the 583−585 nm range, 5 ps duration, 4 MHz repetition rate) were obtained from a synchronously pumped, cavity-dumped dye laser (Coherent 702-2, with rhodamine 6G or pyromethine 567 gain dyes). The dye laser was pumped at 532 nm by the second harmonic of a passively mode-locked, diode-pumped NdIII− YVO4 laser (Spectra-Physics Vanguard). After splitting the total fluorescence emission with a polarizing beamsplitter cube, the parallel (F∥) and perpendicular (F⊥) polarization components were wavelength selected by a pair of double-subtractive monochromators (Spectral Products CM112, 4 nm bandpass) and detected by a pair of microchannel-plate photomultiplier tubes (Hamamatsu R3809U-50). The signals from the two polarizations were accumulated in a two-channel TCSPC system (Becker & Hickl SPC-132). Data acquisition was controlled by LabVIEW routines. For use in the construction of time-resolved spectra, dichroism-free intensity transients at each detection wavelength were calculated from the polarization components as F(t ) = F (t ) + 2F⊥(t )

λ = (ν0 − 0,A − ν0 − 0,F)/2

(4)

−1

is 55 ± 10 cm . This value is more than a factor of 2 smaller than the value of 145 cm−1 obtained previously with the gelfiltration preparations using the same formula,23 but it is comparable to that reported recently for fbCytc in water, 38 cm−1, with preparations isolated using ion-exchange chromatography as described above. These estimates for λ require the assumption that the ground-state and excited-state potentialenergy surfaces are harmonic and have the same normal-mode frequencies,63,64 which is the case if the absorption and fluorescence spectra exhibit an exact mirror symmetry with respect to frequency, and that the vibronic line shapes are Gaussians.54 Estimates for λ that are about a factor of 2 larger are obtained using the formula determined by Jordanides et al.,54 which makes no assumption about the form of the absorption and fluorescence line shapes. The reorganization energy in fbCytc is somewhat smaller than observed in ZnCytc because part of the solvation response is slow enough to be effectively static compared to the fluorescence emission time scale.27 The unfolding transition for ZnCytc can be monitored by measuring the shift to the blue of the absorption and fluorescence spectra that accompanies the addition of Gdm+ to the solution. The 0−0 and 0−1 peaks in the fluorescence spectrum shift to the blue by 40 cm−1 upon addition of 5 M Gdm+, which results in a fully denatured ensemble. In the transition region, with the assumption that the fluorescence spectrum observed in the presence of a certain Gdm+ concentration is the linear combination of the spectra observed in the folded and unfolded states, in water and in 5 M Gdm+, respectively, the wavenumber of the peak maximum of either the 0−0 or 0−1 peak can be used to determine the fraction of unfolded protein present in the ensemble, f U. Figure 3 shows that ZnCytc exhibits a two-state unfolding response65

(3)

The instrument-response width determined with a scattering solution at the sample’s position was 65 ps fwhm.

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RESULTS Continuous-Wave Fluorescence Spectroscopy. Figure 2 shows the Q-band absorption and fluorescence emission spectra obtained from ZnCytc in water at 22 °C and pH 7.0. The spectra are plotted as dipole strengths, AD(v) = A(v)/v (dashed line) and FD(v) = F(v)/v3 (solid line), respectively, with respect to emission wavenumber v.57,61,62 The ZnCytc

−1 ⎛ ⎛ ΔG Fold + mCGdm+ ⎞⎞ ⎟⎟ fU = ⎜1 + exp⎜ ⎝ ⎠⎠ RT ⎝

(5)

as defined by ΔGFold, the Gibbs free energy for folding in water, and m, the rate of change of the Gibbs free energy with respect to CGdm+, the Gdm+ concentration. A fit of the titration profile in Figure 3 to eq 5 obtains the midpoint of the unfolding transition, 1.75 M, and ΔGFold = −10.74 kJ/mol. Similar parameters are obtained from a Gdm+ titration of the absorption spectrum (data not shown). These results show that ZnCytc is significantly less stable than FeIICytc (ΔGFold = −30.9 kJ/mol) under similar conditions.66 Picosecond Time-Resolved Fluorescence Spectroscopy. The FSS response from ZnCytc solutions at 22 °C was determined using a set of picosecond fluorescence intensity transients acquired with 5 nm steps of the emission monochromators from 585−680 nm, across the full width of the 0−1 fluorescence peak from the maximum of the 0−0 peak. Owing to the 5 nm bandpass of the emission monochromator and the relatively small value for the solvation reorganization

Figure 2. Absorption (Q-band) and fluorescence spectra (with excitation at 584 nm) from ZnCytc in water at 22 °C and pH 7.0. The spectra are plotted as absorption and fluorescence dipole strengths, AD(v) = A(v)/v (dashed line) and FD(v) = F(v)/v3 (solid line), respectively, with respect to emission wavenumber v.57,61,62 The spectra are scaled arbitarily so that the 0−0 peaks have the same relative intensity. 14591



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response from fbCytc27 because it has proven in practice to be a more robust measurement and relatively insensitive to the signal/noise ratio for the measured fluorescence transients. Figure 4 compares the FSS response from ZnCytc in water and in 0.25 M Gdm + , the lowest concentration we

Figure 3. Unfolding of ZnCytc at 22 °C and pH 7.0 in the presence of Gdm+, as detected using the wavenumber of the peak maximum of the 0−0 (●, solid curve) and 0−1 (○, dashed curve) fluorescence peaks (spectra not shown). The data points are superimposed upon separate models for a two-state unfolding transition (eq 5). From the average of the two models, the midpoint of the transition occurs at 1.75 M, ΔGFold = −10.74 kJ/mol, and m = 6.16 kJ/mol M.

energy, we were unable to obtain high signal/noise ratios much further to the blue than the peak maximum of the 0−0 peak because of scattering from the excitation laser. The laser was tuned to the wavenumber at which the 0−0 peaks in the absorption and fluorescence dipole-strength spectra cross for a given sample (∼17050 cm−1 in water). This tuning prepares the S1 state of the ZnII porphyrin in the absence of excess vibrational excitation.58,67 The integral of each transient with respect to the time axis was intensity normalized by the integrated fluorescence dipole strength at its emission wavenumber, as determined from the continuous-wave fluorescence spectrum. The full set of normalized transients was then used without deconvolution of the instrument response function to assemble a time−emission wavenumber−intensity (dipole strength) surface, FD(v, t) = F(v, t)/v3. Time-resolved fluorescence spectra were obtained as slices of the surface at each delay point. We excluded from subsequent analysis the spectra falling prior to the 100 ps delay point past the center of the peak of the 65 ps instrument-response profile, as determined using a scattering solution at the sample position. The FSS response function from ZnCytc in water and in the presence of Gdm+ was determined as the time dependence of the mean fluorescence frequency, ⟨v(t)⟩, of the 0−1 peak, which was obtained from a first-moment integration with respect to v over the time-resolved fluorescence dipole-strength spectra, FD(v, t), as a function of the time t

Figure 4. Fluorescence Stokes shift response functions for ZnCytc at 22 °C in water (top panel) and in 0.25 M Gdm+ (bottom panel). The data points were determined as the ⟨v(t)⟩ response obtained as the first moment (eq 6) of the 0−1 fluorescence peak at each time delay point in the fluorescence dipole-strength surface, FD(v, t). Fitted biexponential models (eq 7) are superimposed on the data points; the fit parameters are discussed in the text and are included in Figures 7 and 8.

characterized. In both solutions, the response over the 100 ps to 10 ns regime is well described by a biexponential relaxation function ⟨ν(t )⟩ = A1 exp( −t /τ1) + A 2 exp(−t /τ2) + ν∞

The integration end points for the first-moment calculation of ⟨v(t)⟩, v1 and v2 in eq 6, were set to enclose most of the area of the 0−1 peak but to avoid the spectral congestion and poorly resolved vibronic structure in the spectral region between the 0−0 and 0−1 peaks (see Figure 2); in water, v1 = 15000 cm−1 and v2 = 16100 cm−1. We varied these choices, especially by adding a few additional intensity samples in the congested region to the blue of the 0−1 peak maximum, to determine how the response returned by the integrals in eq 6 was affected. The shape of the FSS response we obtained is relatively insensitive to changes in the integration end points. The FSS response obtained for ZnCytc in water (Figure 4, top panel) is very similar to that reported previously;23 there are two decay (red-shifting) components, with time constants τ1 = 135 ± 36 ps and τ2 = 1.60 ± 0.22 ns, and the faster of the two components exhibits about one-half the amplitude of the

ν

ν(t ) =

∫ν 2 dν νFD(ν , t ) 1

ν

∫ν 2 dν FD(ν , t ) 1

(7)

(6)

This calculation obtains the time-evolving part of the numerator of Sv(t) (eq 1).25 In some of the previous work in this laboratory on ZnCytc’s FSS response,23,47 the FSS response was estimated using the time dependence of the 0−0 transition frequency, which was obtained from a fitted vibronic progression over the 0−0 and 0−1 peaks with a fixed mode frequency. This analysis showed that the spacing of the 0−0 and 0−1 peaks does not evolve with emission time during the FSS response. We used the non-parametric, first-moment approach here and in the more recent work on the FSS 14592



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slower component, A1 = 9.8 ± 1.7 cm−1 and A2 = 20.7 ± 1.4 cm−1, respectively. (The FSS response parameters reported in this paper were obtained as the average and standard deviations of the fit parameters obtained from a set of usually three replicate trials with fresh samples.) The total shift, A1 + A2, accounts for about one-half of the solvation reorganization energy estimated from the continuous-wave fluorescence spectrum, so a significant fraction of the solvation response occurs in the 2 ns time scale where the signal/ noise ratio decreases significantly. The skew (asymmetry) parameter appears to decrease significantly with time, from 1.31 at t = 100 ps to 1.06 at t = 9 ns, but this parameter is poorly determined owing to covariance with the spectral line width, which increases by ∼8% over the same time range. Since these changes occur primarily over the >2 ns range where the signal/ noise ratio is poorer, we suggest that the observed shift can primarily be assigned to dynamic solvation. The time-resolved fluorescence spectra observed from ZnCytc in the presence of 0.25−5.0 M Gdm+ are also effectively of constant line shape over the 100 ps to 10 ns emission time scale. Figure 6 and Table 2 show the timeresolved spectra and line shape parameters we observed for the example of 2.0 M Gdm+, just past the midpoint of the unfolding transition. The fitted line shape of the 0−1 peak is apparently about 5% narrower than observed in water but is comparably skewed. During the time range where the mean frequency, ⟨v(t)⟩, exhibits a bidirectional shift (see Figure 4, bottom panel, for the 0.25 M Gdm+ response), the line shape parameters are essentially invariant. Gdm+ Dependence of the FSS Response. Figure 7 shows a plot of the time constants for the biexponential FSS

Figure 5. Time-resolved fluorescence dipole-strength spectra showing the time evolution of the 0−1 peak from ZnCytc in water at delay times of 100 ps (●), 1 ns (○), 2 ns (◆), 5 ns (□), and 9 ns (▲). The data points in each spectrum are superimposed with log-normal line shapes fitted to the data points acquired over the 14900−16250 cm−1 region. The fit parameters are listed in Table 1. 14593



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responses observed in ZnCytc solutions at pH 7.0 as Gdm+ is titrated across the unfolding transition, from 0 to 5 M. As noted above in the comparison of the FSS responses in water and in 0.25 M Gdm+ (Figure 4), the time constant of the faster of the two response components, τ1, lengthens abruptly when Gdm+ is added to the solution, and the first component is converted from a red shift to a blue shift. The approach to the unfolding transition is marked by a decreasing trend in τ1 from 820 to 440 ps that terminates at 3 M Gdm+, in the presence of a fully unfolded ensemble, before assuming an increasing trend at higher Gdm+ concentrations. In contrast, the time constant for the slower, red-shifting component, τ2, rises from 3.5 ns at 0.25 M to 4.4 ns at 1.0 M, where it appears to saturate. Thus, the τ2 response saturates well below the unfolding midpoint (1.75 M) even though the viscosity of the solution rises rapidly as the Gdm+ concentration increases.72 A similar saturation at low Gdm+ concentrations is inferred from the plots shown in Figure 8 of the amplitudes for the fast

Figure 6. Time-resolved fluorescence dipole-strength spectra showing the 0−1 peak from ZnCytc in 2.0 M Gdm+ at delay times of 200 ps (●), 500 ps (○), 1.2 ns (◆), 4 ns (□), and 8 ns (■). The data points in each spectrum are superimposed with log-normal line shapes fitted over the 14900−16250 cm−1 region. The fit parameters are listed in Table 2.

Table 2. Log-Normal Line Shape Parametersa for the 0−1 Peak of the Time-Resolved Fluorescence Spectra from ZnCytc in 2.0 M Gdm+ at 22° C Shown in Figure 5 delay (ps)

A

v0 (cm−1)

σ (cm−1)

ρ

200 500 1200 4000 8000

1 0.80 0.46 0.19 0.08

15672 15683 15679 15675 15657

590 610 590 590 589

1.31 1.31 1.31 1.31 1.30

a Amplitude (normalized integrated dipole strength), A; center frequency, v0 ; width, σ; asymmetry (skew), ρ.68

Figure 8. Fluorescence Stokes shift response amplitudes for ZnCytc at 22 °C and pH 7.0 as a function of the Gdm+ concentration, as obtained by fitting a biexponential model (eq 7) to the FSS response obtained from the first moment of the 0−1 fluorescence peak (see Figure 6 for examples). The amplitude A1 (top panel) is that for the faster of the two components; negative values represent the shifts to higher frequency (blue shifts), which were observed only in the presence of Gdm+. A2 (bottom panel) is the amplitude for the slower, red-shifting component.

and slow components of the FSS response, A1 and A2, respectively. Aside from a reproducible discontinuity at 3 M Gdm+, at the end of the unfolding transition, both amplitudes exhibit very little change at Gdm+ concentrations above 0.5 M. These results indicate that the overall character of the FSS response from ZnCytc is determined by changes in the hydration shell that occur when Gdm+ is added to the solution but before the average stucture of the protein is significantly altered.

Figure 7. Fluorescence Stokes shift response time constants for ZnCytc at 22 °C and pH 7.0 as a function of the Gdm+ concentration, as obtained by fitting a biexponential model (eq 7) to the FSS response obtained from the first moment of the 0−1 fluorescence peak (see Figure 6 for examples). The time constant τ1 (top panel) corresponds to the faster of the two response components, which is the blue shift observed in the presence of Gdm+; τ2 (bottom panel) is the time constant for the slower, red-shifting component.

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DISCUSSION The transformation of the FSS response from ZnCytc in the presence of Gdm+ shows that the intrinsic ZnII porphyrin senses both protein and hydration-shell motions over the 100 14594



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ps to 10 ns time scale. Even at low Gdm+ concentrations and well below the unfolding transition, the FSS response is transformed from a monotonic red shift of the time-resolved fluorescence spectrum in water to a bidirectional shift with slower time constants. This response is remarkable because it is quite different from that normally associated with a molecular liquid.49 Because the average protein structure around the ZnIIporphyrin probe is essentially unchanged until the unfolding transition region is reached, above 1 M Gdm+, the transformation of the FSS response at lower Gdm+ concentrations chiefly involves changes in the structure and energetics of the hydration shell or of its interface with the protein. These changes are likely to be caused by binding of Gdm+ ions to the protein surface, especially considering that the shape of the plot of the slower, red-shifting time scale (τ2) as a function of the Gdm+ concentration is reminiscent of a ligand-binding saturation curve (Figure 7, bottom panel). Figure 9 describes the diffusive FSS response of ZnCytc and its hydration shell using displaced potential energy curves

transition of the probe is followed by a monotonic FSS response. As the system moves from left to right along the solvation coordinate, the energy gap between the excited-state and ground-state potentials decreases, and the time-resolved fluorescence spectrum shifts to the red. This response is characteristic of a reorganizational response on a potential where only a single, global minimum is thermally accessible.49 The blue-shifting initial portion of the bidirectional FSS response observed from ZnCytc in the presence of Gdm+ directly reports that the protein/solvent system moves along a region of the solvation coordinate where the ground-to-excitedstate energy gap increases. At the same time, because the associated time scales are much longer than associated normally with vibrational equilibration (> ∼10 ps), the response must be purely dissipative; the system moves monotonically on the excited-state potential from high energy to low energy or over low (Ea ≤ kBT) barriers while remaining close to thermal equilibrium. These requirements constrain the structural nature of the FSS response in the presence of Gdm+ to the situation depicted semiquantitatively in Figure 9: a light-induced conformational change of the protein/solvent surroundings of the ZnII porphyrin. Figure 9 shows a set of displaced protein/hydration-shell potentials having two conformational minima separated by a low barrier. In the presence of Gdm+, the excited-state potential includes an alternative conformation that is more stable than that present at equilibrium in the ground state. Even with a small displacement of the excited-state potential, The FSS response expected from the scheme shown in Figure 9 should have three phases. Just as in a simple, single-minimum system, the initial propagation along the excited-state surface from the Franck−Condon region (vertical from the equilibrium groundstate conformation) is associated with a narrowing of the energy gap to the ground state, so a red shift of the fluorescence spectrum should be observed as the first part of the FSS response. This portion would include the 100 ps observation window in the present TCSPC experiment.23,54 Then, as the system moves over a low barrier to the alternative, lower-energy conformation, the energy gap increases; once the barrier is crossed, the energy gap again narrows as the system descends to the minimum of the alternate conformation. It is evident that the blue- and red-shifting second and third parts of the FSS response are directly observed on the >100 ps time scale in the presence of Gdm+. The conclusion that the energy of the alternative, lightinduced conformation in ZnCytc is lower in energy in the excited state than that of the initial, Franck−Condon conformation allows the FSS response to be purely dissipative and remain close to thermal equilibrium. This restriction requires, then, that the relative energy of the two conformations be reversed in the ground state, as indicated schematically in Figure 9. Thus, excitation of the ZnII porphryin is followed by reorganizational motions of the protein and solvent that result in an excited-state change in conformation; upon relaxation to the ground state, owing to emission of a fluorescence photon or non-radiative decay, a reverse conformational change occurs that yields the initial, ground-state conformation. Note that if the energy of the alternative conformation is raised in the absence of Gdm+ so that it is thermally inaccessible in the excited state, the reorganizational response is restricted to the minimum of the initial conformation and

Figure 9. Top panel: Protein/hydration-shell potential-energy curves for ZnCytc in water (dashed curves) and in the presence of Gdm+ (solid curves) with respect to a generalized, one-dimensional solvation coordinate arising from two conformational states of the protein and its coupled hydration shell. The vertically displaced sets of potentials correspond to those accompanying the ground state (Eg) and firstexcited state (Ee) of the ZnII porphyrin. Potential-energy minima for the ground state (xg) and the excited states in water (xe,H2O) and in Gdm+ (xe,Gdm+) are marked on the abscissa and by vertical dotted lines. Bottom panel: Energy gap (ΔE = Ee − Eg) between the excited state and the ground state.

drawn as a function of an effectively one-dimensional coupled protein−hydration-shell solvation coordinate. The two potentials correspond to the ground state and first-excited state of the ZnII-porphyrin. The excited-state potential is displaced along the solvation coordinate with respect to the ground-state potential owing to polar and non-polar factors: the ground-toexcited-state change in permanent dipole moment of the ZnII porphyrin and its molecular size and/or polarizability, respectively. Because the change in dipole moment is small for the ZnII porphyrin in ZnCytc, the non-polar part of the response is likely to be significant. In water, the optical 14595



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hydration-shell water molecules that result in a lengthened solvation correlation time and large reorganization energies owing to thermally promoted transitions between hydrogenbonding states.35 The structural character of the motions that contribute to the FSS response of ZnCytc is not well-defined by the present results, but the ps−ns correlation time scales suggest an intermediate character that includes local and global (at least on the length scale of the ZnII porphyrin) fluctuations. Because the FSS response saturates in the presence of Gdm+ signficantly below the midpoint of the two-state unfolding transition at 1.75 M and apparently remains the same in the presence of the unfolded state, despite its probable heterogeneity, the enhanced protein−hydration-shell fluctuations sensed on the ns time scale by the ZnII porphyrin in ZnCytc are established well prior to the unfolding transition and are probably distinct from the specific motions that directly promote formation of the unfolding transition state. This finding is consistent with the conclusion by Englander and co-workers that the partially unfolded protein regions (foldons) that contribute to the unfolding of cytochrome c in the presence of Gdm+ are not significantly populated prior to the unfolding transition.44,75 Thus, we suggest that the unfolding transition state is stabilized by the Gdm+-perturbed hydration-shell structure so that a distinct class of protein fluctuations promotes crossings to the unfolded product state. These fluctuations are not necessarily confined to the fluorescence time scale sensed by the ZnII porphyrin in ZnCytc, and they might very well be expected to be more internal in character than those that would be directly slaved to motions of the hydration shell.38

results in a monotonic, red-shifting FSS response. (In Figure 9, the excited-state and ground-state energies for the alternate configuration are raised by an equivalent amount upon moving from Gdm+ to water.) The results indicate, then, that the alternative conformation is stabilized in the presence of Gdm+ even at concentrations that are much lower than required to denature the protein. The situation described by Figure 9 is consistent with a lightinduced change in conformation of the ZnCytc protein and its hydration shell in the presence of Gdm+ that results in an expanded cavity around the excited-state ZnII porphyrin. Expansion of the cavity, or an equivalent reduction in the surrounding density (or polarizability), would effectively describe a non-polar solvation response to the light-induced change in molecular size of the ZnII porphyrin.56,73 In the absence of Gdm+, however, the protein and solvent structure is stabilized (and/or the barrier to the expanded conformation is raised) enough to maintain the original conformation so that a conventional red-shifting FSS response is observed. This consideration argues against a structural assignment of the cavity expansion to the reorganizational response that accompanies photodissociation of the axial ligands to the ZnII ion, the side chains of the methionine (M80) and histidine (H18) residues (see Figure 1).43 As shown in a previous study,47 the axial-ligand response is readily observed in water in terms of a change in the relative intensities of the 0−0 and 0−1 fluorescence peaks that occurs on different time scales from the shifts of the time-resolved fluorescence spectrum that are characterized in the FSS response. In terms of conformational forces, the addition of Gdm+ increases the f lexibility of the protein and solvent that surrounds the ZnII porphyrin: the potential is effectively softened (the restoring force decreases), so the system makes a larger displacement along the solvation coordinate than in the absence of Gdm+. This conclusion is consistent with the change in correlation time scales detected in the FSS response. For Brownian diffusion on an effectively one-dimensional energy landscape, the correlation times are proportional to the meansquared deviation for the characteristic fluctuation and inversely proportional to the friction, which relates to the characteristic modulation or roughness of the energy landscape in the potential-energy minimum being sampled.27 Thus, the 4.4 ns time constant observed in Gdm+ for the red-shifting segment of the response is consistent with fluctuations having 3 times larger amplitudes than in water, where a 1.6 ns time constant is observed. This interpretation depends on the assumption that the character of the coupled protein and solvent motions detected in the FSS response are comparable for ZnCytc in water and in low concentrations of Gdm+. Changes in the hydrogen-bonding structure of water occur in the presence of Gdm+,42 and the overall viscosity of the hydration-shell medium would be expected to increase rapidly as Gdm+ is incorporated,72 but the time scales detected in the FSS response depend on the Gdm+ concentration in a manner that is reminiscent of a binding equilibrium (see Figure 7). This observation favors the conclusion that the action of Gdm+ involves binding to sites on the protein surface that are accessible to water molecules in the hydration shell.41,74 Accordingly, the simulations by Friesen and Matyushov of the electric-field fluctuations in hydration shells near surface dipoles or charges anticipate several aspects of the response observed in the present study. The binding of ions like Gdm+ at surface sites will produce defects in the network of the adjacent

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CONCLUSIONS The ps−ns FSS response of ZnCytc is transformed in the presence of low concentrations of Gdm+ from the monotonic red-shifting response that is typical of molecular liquids and of ZnCytc under native-state conditions to a distinctive bidirectional response that begins with a shift of the time-resolved fluorescence spectrum to the blue. This response is characteristic of a diffusive search of a coupled protein−hydration-shell potential surface having at least two conformational minima. The blue-shifting part of the FSS response accompanies a lightinduced change in conformation of the protein and solvent that expands the cavity surrounding the ZnII porphyrin. Because the bidirectional FSS response is observed at low Gdm + concentrations and well below the unfolding transition, its origin is a perturbed hydrogen-bonding structure for the hydration shell caused by binding of Gdm+ to the protein surface. The protein/solvent surroundings of the Zn II porphyrin are rendered more flexible by the changes in the hydration-shell structure so that the amplitudes and correlation times for the characteristic fluctuations that contribute to the detected solvation response are markedly increased.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Present Addresses †

Department of Physics, School of Applied Science, KIIT University, Bhubaneswar, Orissa, India 751024. ‡ Naval Medical Research Center, 503 Robert Grant Ave, Silver Spring, MD 20910. 14596



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Department of Chemistry, University of Chicago, 575 S. Ellis Avenue, Chicago, Illinois 60637. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was supported by the National Science Foundation Biomolecular Systems Cluster/Molecular Biophysics program under grant MCB-0920101. We thank Professor Gary Blanchard (Michigan State University) for the use of the picosecond fluorescence spectrometer in his laboratory and for his technical assistance with the experiments. We also thank Professor Sergei Savikhin (Purdue University) for comments on the results. J.J.M. was supported by the United States Navy in a Duty-Under-Instruction status. Her contributions to this work were performed as a part of her official duties and as part of her work towards a Ph.D. in Chemistry at Michigan State University. Disclaimer: The views expressed in this publication are those of the authors and do not necessarily reflect the official policy or position of the Department of the Navy, Department of Defense, or the United States government.

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Dynamic Solvation and Coupling of the Hydration Shell of ZnII

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