Broad-Band PumpâProbe Spectroscopy Quantifies Ultrafast Solvation

Letter pubs.acs.org/JPCL

Broad-Band Pump−Probe Spectroscopy Quantifies Ultrafast Solvation Dynamics of Proteins and Molecules Chanelle C. Jumper,† Paul C. Arpin,†,‡ Daniel B. Turner,†,§ Scott D. McClure,† Shahnawaz Rafiq,∥ Jacob C. Dean,∥ Jeffrey A. Cina,⊥ Philip A. Kovac,⊥ Tihana Mirkovic,† and Gregory D. Scholes*,†,∥ †

Department of Chemistry, University of Toronto, 80 St. George Street, Toronto, Ontario M5S 3H6, Canada Department of Physics, California State University, Chico, Chico, California 95929-0202, United States § Department of Chemistry, New York University, 100 Washington Square East, New York, New York 10003, United States ∥ Department of Chemistry, Princeton University, Washington Road, Princeton, New Jersey 08544, United States ⊥ Department of Chemistry and Biochemistry, and Oregon Center for Optical, Molecular, and Quantum Science, University of Oregon, Eugene, Oregon 97403, United States

J. Phys. Chem. Lett. 2016.7:4722-4731. Downloaded from pubs.acs.org by UNIV OF SOUTH DAKOTA on 09/18/18. For personal use only.

‡

S Supporting Information *

ABSTRACT: In this work, we demonstrate the use of broadband pump−probe spectroscopy to measure femtosecond solvation dynamics. We report studies of a rhodamine dye in methanol and cryptophyte algae light-harvesting proteins in aqueous suspension. Broad-band impulsive excitation generates a vibrational wavepacket that oscillates on the excited-state potential energy surface, destructively interfering with itself at the minimum of the surface. This destructive interference gives rise to a node at a certain probe wavelength that varies with time. This reveals the Gibbs free-energy changes of the excited-state potential energy surface, which equates to the solvation time correlation function. This method captures the inertial solvent response of water (∼40 fs) and the bimodal inertial response of methanol (∼40 and ∼150 fs) and reveals how protein-buried chromophores are sensitive to the solvent dynamics inside and outside of the protein environment.

T

effect),9,11−13 and three-pulse stimulated photon echo peak shift spectroscopy (3PEPS).14−17 Theoretical considerations and femtosecond pulse experiments facilitated the discovery of an initial Gaussian response that makes up a large proportion of equilibration on an ultrafast time scale, ∼100 fs or faster followed by slower diffusive picosecond components.18−24 The inertial component can be particularly challenging to capture because of these fast dynamics, and it is limited by the duration of an exciting pulse and by the time resolution of the given experiment. Fluorescence up-conversion and 3PEPS measurements have provided the most insight into the inertial response.3 However, the former has not been reported with a time resolution faster than 40 fs,25 and the latter is generally more easily analyzed for population times longer than 50−100 fs due to obstruction by other signals at short times,26 unless other complicated fitting procedures,15,27−30 data processing,31−33 or additional experimental approaches31 are invoked. As a result, the shortest time response reported has decreased as a function of experimental

he Stokes shift is a measure of the difference between the maximum energies of absorption and fluorescence and is affected by the dielectric properties of the solvent. The Stokes shift can evolve in time after photoexcitation, and this dynamic response to charge redistribution upon light absorption of solute molecules occurs over a wide range of time scales, from femtoseconds to nanoseconds. The solvation correlation function represents the time development of the Stokes shift and is a key parameter for the investigation of dynamical processes in the condensed phase. The solvation spectral density is related to the solvation correlation function C(t) through the fluctuation−dissipation theorem.1−3 Contributions to C(t) (eq 1) may include both intramolecular vibrational motions and system−bath energy dissipation, where E is the free energy of solvation at times t, 0, and infinity C(t ) =

E (t ) − E (∞ ) E(0) − E(∞)

(1)

The evolution of the dynamic Stokes shift that represents the excited-state free-energy equilibration generally occurs over several time scales that can be monitored by different spectroscopic methods including time-dependent fluorescence spectroscopy,4 fluorescence up-conversion spectroscopy,5,6 transient grating,7−10 transient birefringence (optical Kerr © 2016 American Chemical Society

Received: September 29, 2016 Accepted: November 4, 2016 Published: November 4, 2016 4722

DOI: 10.1021/acs.jpclett.6b02237 J. Phys. Chem. Lett. 2016, 7, 4722−4731

Letter

The Journal of Physical Chemistry Letters advances. In all cases, finite pulse durations limit the extraction of the initial time dependence of the correlation function. Here we use coherent wavepacket evolution analysis34−36 to resolve the ultrafast component of the solvation response for a model molecular system in organic solvent and a set of aqueous chromophore−protein systems. Transient absorption spectra of molecules in solution may reveal oscillatory modulations that are typically signatures of vibrational wavepackets prepared by the ultrashort excitation (pump) pulse.37,38 Upon femtosecond photoexcitation of organic molecules, Franck−Condon-active molecular vibrational wavepackets can be launched on the ground- and excited-state potential energy surfaces. The distribution of vibrational amplitudes between the ground and excited states depends on the duration,39 chirp,40 and spectrum of the pump pulse.34−36,41−43 Excited-state vibrations can be selected by using a sufficiently brief pulse with a spectrum that covers the entire absorption spectrum of the sample, inhibiting impulsive-stimulated Raman excitation of coherent nuclear motion in the electronic ground state. A node present in the amplitude profile of excited-state oscillations occurs at the minimum of the potential energy surface due to the interference of antiphasing of oscillations on either side of the peak emission frequency.44 Figure 1 illustrates the evolution of the node over time on the excited-state potential energy surface as probed by broad-

band transient absorption spectroscopy. In the interpretation of the time-resolved dynamics of this node, it is useful to separate the free energy picture into orthogonal contributions from intramolecular and intermolecular degrees of freedom (as plotted in Figure 1: nuclear coordinate and bath coordinate).45−49 The impulsive excitation of a molecule reaches a Franck−Condon state by vertical transition. Vibrational relaxation from higher vibrational states involves dissipating excess energy through vibrational cooling-transfer processes that involve transfer of energy to the bath and intramolecular vibrational relaxation. With a broad-band probe, stimulated emission is measured at a range of probe wavelengths with superimposed oscillations of the Franck−Condon-active modes. The net oscillating component of the pump−probe signal carries an opposite phase on either side of the peak emission frequency. A node in the stimulated emission signal is caused by the destructive interference of intramolecular vibrational modes of the molecule at the minimum of the multidimensional potential energy surface where destructive interference is at a maximum. Because the nuclear displacement of all of the intramolecular modes is constant with respect to the ground-state potential, the position of maximum destructive interference does not change as a function of vibrational relaxation. Thus, experimentally, there is a single node that captures the constant multidimensional interference of all vibrational modes involved in the electronic transition and its position is insensitive to dynamics from vibrational cooling and intramolecular vibrational redistribution (IVR). However, assuming that the molecule is linearly coupled to the solvent, there exists a quadratic free-energy relationship of the excited state with respect to the bath coordinate. Solvent relaxation lowers the energy of the excited-state potential, shifting the emitted photon energy to lower energy values. As a result, the peak emission frequency and node position shift to lower energies as a function of only solvent relaxation.50 Figure 1 illustrates how the minimum of the excited-state potential energy surface is constant along the nuclear coordinate for a single nuclear harmonic potential, while the free energy of the entire surface changes as a function of the evolving bath coordinate. Overall, the intramolecular oscillations are a probe of intermolecular (solvent reorganization) dynamics, and our goal is to investigate the use of this method for resolving the ultrafast solvation response. The benefit of using wavepackets to resolve the solvation response is that the node position is not dependent on population dynamics or convolution with other signals. This method also allows us to investigate more complex (multichromophoric) systems that would otherwise be complicated by energy transfer. In this work, we use these excited-state vibrational signals to extract the solvation dynamics for rhodamine 640 perchlorate in methanol and four phycobiliproteins in water.51,52 We compare our results to values obtained from established methods of resolving the solvation dynamics. These samples were chosen in order to explore different examples of solvation dynamics that we can measure with this method. The solvation response of aqueous macromolecular systems such as the protein−chromophore environment is of particular interest because the inertial response is not completely understood.53−56 It is not clear whether the protein or the solvent dominates the ultrafast response57−59 or has functional effects on important biological processes such as enzyme

Figure 1. Schematic illustration of nested free-energy curves involved in solvation dynamics after excitation (red) along orthogonal degrees of freedom from a generalized bath coordinate (black) and nuclear coordinate (blue). The node position is the minimum of the multidimensional excited-state potential energy surface in the nuclear coordinate where destructive interference in the wavepacket is at a maximum. The free energy of the surface (in the nuclear coordinate) is lowered as a function of linear coupling to the solvent, and the wavepacket travels along the quadratic free-energy curve in the generalized bath coordinate. The time evolution of the node represents dissipation of the excited-state energy due to the collective response of the bath motions (inertial and diffusive). In accordance with a one-dimensional displaced harmonic oscillator model coupled to a bath, the position of the node at the minimum of the harmonic well implies that free-energy changes as a result of intramolecular relaxation are not captured, and this method primarily isolates the response of the bath. 4723


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The Journal of Physical Chemistry Letters activity, electronic energy transfer,60 or intraprotein electron transfer.61 On the basis of our results, we discuss some of the general features associated with a protein−chromophore environment and factors associated with the assignment of the inertial response. Molecular Solvation Dynamics. The solvation dynamics of methanol were examined by using rhodamine 640 perchlorate as a probe molecule. Rhodamine dyes belong to the class of xanthene dyes with the transition moment parallel to the long axis of the molecule.62 A carboxyphenyl group attached to the central carbon in xanthene causes molecules of rhodamine 640 (and similarly of rhodamine B) to be found in one of three main molecular forms: cation, lactone, or zwitterion.63−65 Rhodamine 640 (also known as rhodamine 101) takes the zwitterionic form in dilute polar protic solutions of the perchlorate, and the molecular form can be assigned based on the absorption and fluorescence maxima. Owing to inductive effects, the zwitterion exhibits a hypsochromic shift in polar solvents compared to the cationic or lactone form.63−65 Rhodamine 640 perchlorate steady-state spectra are shown in Figure 2a with an absorption maximum of 567 nm (17600 cm−1) and a fluorescence maximum of 590 nm (16900 cm−1), characteristic of the zwitterionic form of the molecule in methanol. This form of rhodamine dye (Rh B, Rh 640/Rh 101) exhibits moderate solvatochromism66 and a significant increase in the Stokes shift in methanol compared to that in gas-phase measurements for similar rhodamine dyes.67 The change in dipole moment between ground and excited states for the related dye rhodamine B has been calculated from solvatochromic experiments to be 5.33 D.68 In our ultrafast time-resolved measurement, the same broadband pulse was used for both the pump and the probe, shown in red in Figure 2a. The position of the absorption and fluorescence maxima of the dye and the moderate Stokes shift that can be fully captured within the pulse frequency width makes this dye suitable as a probe for these experiments. The pulse duration at fwhm was approximately 14 fs, determined by a polarization-gating frequency-resolved optical gating measurement (PG-FROG). Figure 2b displays residuals of the pump−probe spectra for rhodamine 640 in methanol, after the population dynamics were subtracted. The population decays were fitted by a biexponential and subtracted at each probe wavelength, and high-frequency modes (1000 cm−1 methanol oscillation) were filtered out in the frequency domain with a Super Gaussian filter.35,36 The methanol oscillation is a contaminating nonresonant solvent signal from impulsive Raman scattering. The data filtration procedure along with oscillation maps before and after removal are shown in Supporting Information Figure S1. A dynamically shifting node is visible in the data (Figure 2b), beginning near the absorption maximum and approaching the fluorescence maximum. The time evolution of the probe frequency at the node was used to generate a correlation function for the solvent response according to eq 1. The probe frequency position of the node was determined according to the position of minimum amplitude in the coherence spectra. Figure 2c plots the resulting correlation function, fitted by a biGaussian plus exponential decay (refer to the discussion below). The resulting fit parameters are listed in Table 1. The two central half-widths of the bi-Gaussian fit are 37 and 144 fs followed by a ∼700 fs exponential decay. The excited-state oscillations are damped before longer time dynamics can be captured (tens of ps).

Figure 2. Steady-state and dynamic spectroscopy of rhodamine 640 perchlorate in methanol. (a) Absorption (black) and fluorescence (blue) and pulse spectrum (red) used for transient absorption measurements. (b) Residual transient absorption spectrum for rhodamine 640 in methanol after subtraction of biexponential population decays determined at each probe wavelength and frequency domain filtration of high-frequency solvent modes. A red-shifting node can be seen in the oscillatory features, which approaches the fluorescence maximum. (c) Correlation function generated by eq 1 using the average node frequency and fit by the sum of a bi-Gaussian and exponential decay. Five independent data sets were used to determine the average node value, and error bars represent the standard deviation.

The reorganization energy listed in Table 1 is estimated from the maxima of the absorption and fluorescence energies, which does not distinguish between contributions from intramolecular and intermolecular sources of energy dissipation. The Stokes shift for a set of rhodamine dyes decreases significantly in the absence of solvent according to gas-phase measurements, and we estimate that between 1/3 and 2/3 of the reorganization energy is due to solvent effects.67 In polar solvation dynamics, inertial solvent motions are responsible for a large proportion of the solvation response occurring on an ultrafast time scale.18,19,69−71 This motion has been referred to as “free-streaming” and refers to the initial 4724


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The Journal of Physical Chemistry Letters Table 1. Estimated Solvation Relaxation Times for Rhodamine 640 in Methanola ER −1

688 (cm )

A1

(1/2)Δω1−1

A2

(1/2)Δω2−1

A3

τ3

E

0.35

37.0 fs

0.50

144 fs

0.13

733 fs

2%

The values of the relaxation times and their relative weights are determined by fitting the function C(t) = A1 exp(−(1/2)ω12t2) + A2 exp(−(1/ 2)ω22t2) + A3 exp(−t/τ3) + E to the time dependence of the node. Δω is the full Gaussian width at half-maximum defined by 2(2 ln 2)1/2ω. ER is the reorganization energy estimated by the total Stokes shift taken from the difference in absorption maxima and emission maxima. E is the portion of relaxation that has not been captured on the time scale of the experiment (within 4 ps). a

Figure 3. Dynamic spectroscopy and X-ray crystal structures of phycobiliproteins. Residual transient absorption spectra after subtraction of biexponential population decays determined at each probe wavelength for (a) PC577, (b) PC612, (c) PC630, and (d) PC645. A red-shifting node can be seen in the oscillatory features, which approaches the fluorescence maximum for each protein. (b) X-ray crystal structures of each protein as labeled. Embedded molecules in red, blue, and yellow represent DBV, PCB, and MBV chromophores, respectively.

predicted for methanol OH libration by molecular dynamics simulations, a ∼30 fs component about Ixx and two degenerate ∼100 fs components about Iyy and Izz.71,74 Therefore, two distinct frequencies are involved in the inertial component as rotations about all three inertial axes cause rotation of the dipole moment responsible for solvation dynamics. The inertial component of methanol solvation was calculated by molecular dynamics simulations and first observed by fluorescence up-conversion by Rosenthal et al.70 The fast component was described by a single-mode Gaussian with a half-width of approximately 100 fs, followed by 560 fs and 8.1 ps exponential decays. With 50 fs resolution, this was the first observation of an ultrafast component in methanol solvation. It

frictionless velocities exhibited by the solvent molecules prior to perturbation of the solute charge distribution (i.e., through excitation).20,72,73 This free-streaming motion at early times after perturbation is sufficient to accomplish much of the solvation response, resulting in an overall Gaussian response function at very short times.18−20 The Maxwell−Boltzmann velocity distribution depends on only the temperature and the inertial properties of the solvent, which are unique for methanol compared to other small-molecule polar solvents such as water and acetonitrile. Methanol has three principal moments of inertia, Ixx, Iyy, and Izz, defined by the x-axis along the C−O bond and the y-axis in the plane defined by the COH group. These inertial moments result in two distinct components 4725


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The Journal of Physical Chemistry Letters is also possible to fit our data by the sum of a single-mode Gaussian and exponentials with Gaussian half-widths of 113 and 720 fs but with a reduced quality of fit. These values are close to the reported values of Rosenthal et al. for methanol for the first two components (Gaussian plus exponential). It is likely that the single-mode Gaussian component of 113 fs represents a weighted average of the two distinct inertial components in methanol, which appear to be resolvable by our method and are best described by a bimodal Gaussian. Other reports for methanol solvation correlation times generally follow multiple exponential decays (or Gaussian plus exponentials) with time constants on the order of 100 fs, 1 ps, and 10 ps.6,15,74−82 To describe the early time dynamics observed by time-resolved fluorescence of coumarin 153, Horng et al. included a 30 fs Gaussian component in their fit, combined with a slower 280 fs response, although the 30 fs was included arbitrarily as the limit of the instrument response function.6 A later report on coumarin 153 in methanol reproduced the 100 fs inertial time scale obtained by Rosenthal et al. (above) but reported an exponential decay for the fit.75 Gumy et al.,80 Zhang et al.,81 and Meulen et al.82 obtained a Gaussian response for the inertial component on the time scale of approximately 100−200 fs. Whether or not a Gaussian component has been included in the literature fits to represent the inertial response generally depends on the time resolution of the experiment. Even when the ultrafast component is generally observable in the 100−300 fs range, the initial curvature of the Gaussian component may still not be resolvable and may only capture the tail end, better fit by an exponential function. Our results resolve the Gaussian component centered at two frequencies described by the differing inertial moments in methanol. Protein Solvation Dynamics. Photosynthetic proteins are good model systems for investigating the aqueous protein solvation response function because they incorporate natural chromophores that serve as probes. Broad-band excitation of phycobiliproteins generates vibrational wavepackets in the excited state that allow us to monitor the solvation dynamics. The crystal structures of the four phycobiliproteins investigated, PC577, PC612, PC630, and PC645, are shown in Figure 3. The absorption and emission spectra for each protein are shown in Figure S2 with the pulse overlap for the broad-band pump− probe experiment. Steady-state spectroscopic parameters for the four proteins are tabulated in Table 2. The absorption and

The resulting background-subtracted pump−probe residuals for each are shown in Figure 3, which reveal oscillations at different frequencies and a zero-amplitude node, shifting toward the red in time. This node very closely approaches the fluorescence maximum for each protein within the time scale of the experiment (4 ps). The frequency and phase of these oscillations have been previously studied, which revealed a phase shift of the oscillations at the node position, indicating the minimum of the excited-state potential energy surface.51,52 Monitoring this nodal position over time reveals the transition frequency correlation function, which within the linear response approximation reveals the Stokes shift response function. The time evolution of the node has been described for a single molecule. 44 For a protein that contains eight chromophores, all potentially experience dynamics from solvent reorientation. Why are there not more nodes from chromophores at different spectral positions? It is possible that spectral offsets between the different chromophores overlap the oscillation maps so as to cancel out the nodal features to the blue and we can only resolve the node relating to wavepackets for the most red-absorbing chromophore (e.g., PCB 82 bilin in PC645).83,84 Furthermore, this is the only chromophore that does not transfer energy rapidly to other chromophores in the complex, a process that decoheres vibrational wavepackets. Second-derivative plots of the oscillatory residuals of PC577 data and simulations, as well as Fourier transform maps in Figure S3, indeed reveal some nodal features at wavelengths further to the blue from the terminal chromophore;44 however, these are not strong enough to resolve Stokes-shifting dynamics from other chromophores in the light-harvesting complex. The probe energy of the red-edge node was measured as a function of time for five independent data sets for each protein. The average correlation function (by eq 2) and standard deviation are plotted in Figure 4. The correlation functions were best fit by the sum of a Gaussian function plus exponential. The fitting parameters are tabulated in Table 3. Comparing data for the four proteins, it appears that the time

Table 2. Phycobiliprotein Steady-State Spectroscopic Parameters in H2O absorption max (blue edge) nm (cm−1) PC577 PC612 PC630 PC645

577 577 583 585

(17300) (17300) (17200) (17100)

absorption max (red edge) nm (cm−1) 606 613 629 645

(16500) (16300) (15900) (15500)

fluorescence max nm (cm−1) 640 641 656 662

(15600) (15600) (15200) (15100)

fluorescence profiles are different for each protein depending on the structure of the protein, the positions of the chromophores, and the couplings between them. The broadband pulse overlaps well with the entire absorption spectrum, thereby exciting each embedded chromophore at time zero and initiating both energy-transfer dynamics as well as solvation dynamics.

Figure 4. Solvation dynamics. Correlation function (according to eq 2) for the four phycobiliproteins, (a) PC577 (b) PC612 (c) PC630, and (d) PC645. Five independent data sets were used to determine the average node value, and error bars represent the standard deviation. The average data are fit by the sum of a Gaussian and exponential decay (red curve). 4726


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The Journal of Physical Chemistry Letters

example, for eosin in the aqueous lysozyme complex87 and equilenin and coumarin 183 bound to the active site of a ketosteroid isomerase protein89 and for the dye DCM in Human Serum Albumin.90 In the ketosteroid isomerase, the active site was interpreted as being an electrostatically rigid environment.89 A similar reduction in the Stokes shift is seen comparing a dye in methanol versus that in a polymer glass at room temperature. 3 The interior of a protein where chromophores bind is generally considered to be hydrophobic, and nonpolar environments are reported to be inefficient at dissipating energy from excited states of chromophores.91 This may limit the overall solvation response. On the basis of these simple arguments of electrostatic rigidity and reduced polarity, it has been argued that dissipation can be directed to water molecules through longer-range interactions.85,87 Even when the protein shields a chromophore from the solvent, the solvent can still dominate the inertial response, though the mechanism is not understood.57,85,92 If inertial protein motions are involved in equilibrating a newly formed electronic charge distribution, then we must consider the motions available to residues on the interior of a protein or active sites, where chromophores and ligands bind. The contribution to the inertial response of small solvent molecules such as water, methanol, and acetonitrile is large, owing to their small inertial moments. The relative amplitude of the inertial response has been shown to decrease as a function of molecular length for a range of alcohols from methanol to decanol6 and for methanol, ethanol, and a series of alkanenitriles.80 It was reasoned that inertial solvation is more efficient in small solvents because inertial motion can involve rotation of the whole solvent molecule, while in longer molecules, contribution is limited to the polar heads. In a protein environment, similar restricted inertial motions of polar or charged side groups may be expected and have been calculated.93 The results in this work reveal that the amplitude of the inertial response is at least 75%, which is characteristic of the inertial component of small-molecule polar solvents. However, proteins can undergo internal motion on femtosecond to picosecond time scales, and fast fluctuations of protein side groups have been thought to contribute to the overall relaxation.7,94 Molecular dynamics simulations have suggested that relaxation and atomic and velocity displacements can occur in proteins on sub-100 fs time scales.95−99 Indeed, it has been shown that a single hydrogen bond between a chromophore and protein environment entirely dominates the solvent response.100 Considering all of these factors combined, solvation response in a chromophore−protein−solvent system is likely as unique as each protein itself, and such properties may even be subject to evolutionary pressure.89 In a review of a large collection of chromophore−protein solvation dynamics, Gilmore argues that under certain conditions the surrounding protein dominates the dynamics of the excited chromophore and in others the solvent dominates.57 The most direct example of a free- versus proteinbound chromophore is the comparison between eosin in solution and that bound to the lysozyme protein by Jordanides et al.87 Here, the mechanism of the ultrafast response was shown to be identical and dominated only by librational motion of bulk water molecules, while longer time scales show deviations owing to the protein response, which can approach milliseconds. At least in small soluble proteins this may be the case.

Table 3. Estimated Solvation Relaxation Times for the Set of Four Phycobiliproteinsa protein

A1

(1/2)Δω1−1 (fs)

A2

τ2 (ps)

PC577 PC612 PC630 PC645

0.752 0.852 0.908 0.892

62.4 54.2 44.2 22.0

0.247 0.148 0.067 0.104

9.20 3.66 0.386 0.305

a

The values of the relaxation times and their relative weights are determined by fitting the data by a Gaussian plus exponential decay function C(t) = A1 exp(−(1/2)ω12t2) + A2 exp(−t/τ2) + E to the time dependence of the node. Δω is the full Gaussian width at halfmaximum defined by 2(2 ln 2) 1/2 ω. At least 97.5% of the reorganization is captured by these biexponential decays (i.e., E ≈ 0).

scales for solvent relaxation depend on the protein structure. The Gaussian response decays with a half-width of 22.0−62.4 fs depending on the protein, which makes up at least 75% of the overall solvent response. The fastest response occurs in the proteins having the closed structure (PC645 and PC630). The second time constant is also variable between the open and closed protein structures. For the open structures PC577 and PC612, the second time constant is on the picosecond scale, ∼9 and ∼4 ps, respectively. For the closed structures PC630 and PC645, the second time constant is 300−400 fs, about an order of magnitude faster than that in the open structures. However, it should be noted that there might be more error associated with the second lifetime due to the time constraints on the experiment. It is possible that either the differing levels of solvent accessibility of the emitting chromophore or the differing local protein environments are responsible for the variability in solvent relaxation times.85,86 A chromophore−protein−solvent system is complicated and exhibits a broad range of energetic couplings and solvation and equilibration time scales.57−59 Upon photoexcitation, three main sources of environmental relaxation may contribute over different time scales: the protein, bulk water, and “biological water”, that is, those that are bound for long time scales on the surface of the protein. Long-range interactions with bulk water are characterized by a dielectric response exhibiting an ultrafast (tens of femtoseconds) component and complete relaxation by tens of picoseconds.16,19,87 Protein reorientation occurs on a nanosecond time scale.88 We can see in these results that over 97% of the equilibration is obtained within the time scales that are equivalent to interaction with bulk water, indicating that slower nanosecond protein reorientation does not contribute significantly. The inertial response in a chromophore−protein−solvent system53 and other aqueous macromolecular systems54−56 is not completely understood. A debated question is whether the protein or the solvent dominates the ultrafast response,57−59 and it is unclear if the inertial response has functional effects on important biological processes like enzyme activity, electronic energy transfer,60 or intraprotein electron transfer.61 We will discuss some of the general features associated with a protein− chromophore environment and factors associated with the assignment of the inertial response: reorganization energy, rigidity of the protein environment, inertial moments, and ultrafast protein side group fluctuations. The magnitude of the reorganization energy reflects the solvation capacity of a given medium. The Stokes shift of a chromophore embedded in a protein has been shown to be smaller than that of the same chromophore in solution, for 4727


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the stored protein samples were thawed and dialyzed against a 0.025 M phosphate buffer (pH7.5) to remove excess ammonium sulfate. Proteins were diluted to an optical density (O.D.) of 0.15/mm. Steady-State Spectroscopy. Solution-phase steady-state measurements were recorded at room temperature. Absorption spectra of rhodamine 640 perchlorate and phycobiliproteins were recorded using a Varian Cary 6000i UV−vis spectrometer with a resolution of 1 nm at room temperature. Fluorescence emission and excitation spectra were recorded on a Fluorolog-3 (Horiba) spectrometer in a 90° mode, with a xenon arc light source and an R928 photomultiplier tube. Pump−Probe Experiments. The transient absorption spectrometer setup is described in detail elsewhere.52 Broad-band pulses were generated from a commercial 5 kHz Ti:sapphire laser amplifier pumped into a home-built noncollinear optical parametric amplifier (NOPA). For rhodamine experiments, the spectrum was tuned centrally to 580 nm and extended past 650 nm, overlapping well with the absorption spectrum (Figure 2). The pulse was compressed with grating and prism compressors to ∼14 fs, as determined by a polarization-gating frequencyresolved optical gating measurement. For protein experiments, the spectrum was tuned centrally to 580 nm with a weak tail extending past 750 nm, overlapping well with the entire protein absorption spectra (Figure S2). The pulse was compressed to ∼10 fs by grating and prism compressors, and the pulse duration was determined by a transient-grating frequencyresolved optical gating measurement. The pulse was split into identical pump, probe, and reference beams with a 1° UV fused silica wedge. The probe beam was directed through the sample and was dispersed and focused onto a 1.25 kHz CCD detector. Rhodamine dyes were measured in a 1 mm glass cuvette. Proteins samples were kept in an ice bath and flowed through a 1 mm cuvette at a rate of 1 mL/min with a peristaltic pump. The sample was pumped at 14 nJ/pulse for proteins and 12 nJ/ pulse for dyes, and successive sets of four probe pulses with the pump blocked and unblocked were recorded. The pump and probe beams had parallel polarization and were focused onto 1/ e beam diameters of 40 and 15 μm, respectively. The probe intensity was balanced by the reference beam intensity as measured on a photodiode to eliminate influences from laser fluctuations during the experiment. The ΔI/I value was then determined by

Our results demonstrate that different protein local environments may at least partially contribute to a change in the inertial response, evidenced by the varying inertial response times of the open vs closed protein structures investigated. This is a complicated topic, and it is clear that more experimental input is needed on the solvation capabilities and polarity of a protein.85 Simple assumptions of a nonpolar homogeneous environment, characterized by a low dielectric constant, do not allow for a complete understanding of potentially finely tuned dynamics occurring within a protein. Finally, the role of solvation relaxation on other functional effects in proteins and biomolecules is not fully understood either. We see in these phycobiliproteins that solvent relaxation can occur on time scales comparable to electronic energy transfer and furthermore that energy transfer is faster in the structures that exhibit the faster solvent relaxation.51 Overall, coherent wavepacket evolution analysis offers an alternative to other methods for determining the early response times of the environment to a molecular charge in solvent or embedded in a protein, especially concerning multiple chromophoric systems. While 3PEPS has successfully been used to monitor solvation response of embedded chromophores in proteins, multiple chromophore systems represent a special case. Capturing the solvation dynamics around photosynthetic proteins is challenging due to complications by energy-transfer events; thus, coherent wavepacket evolution analysis may offer an alternative. When performed with sufficiently short pulses, the source of the signal is from the excited-state vibrations, and we obtain an accurate description of the early time dynamics. Pulse overlap that would normally complicate the extraction of M(t) from 3PEPS signals and even convolution in fluorescence up-conversion101 do not complicate the node, and the excited-state equilibration can be modeled by exponential decays beginning as early as 10 fs. The time evolution of the node is also free of oscillations that are seen in fluorescence up-conversion.70 Overall, this is a promising method to elucidate early time solvation dynamics around a wide range of sample types.

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EXPERIMENTAL SECTION Growth and Isolation of Light-Harvesting Complexes. Hemiselmis pacifica (CCMP706) was cultured in an enriched seawater medium, Prov50 (from NCMA), on a 12/12 h dark−light cycle of 20 μE m−2 s−1 in the growth chamber at 16 °C, while Chroomonas (CCAC 1627 B, also M1627) was grown under identical conditions but in the L1 medium (also from NCMA). Chroomonas mesostigmatica (CCMP269) was cultured at room temperature in either K or Prov50 media from NCMA on a 12/ 12 h dark−light cycle of 20 μE m−2 s−1. Cells were harvested by centrifugation and were resuspended in a 0.1 M sodium phosphate buffer (pH 7.5). Phycocyanin 577 (PC577) was isolated from Hemiselmis pacifica (CCMP706), phycocyanin 630 (PC630) was obtained from Chroomonas (CCAC 1627 B, also M1627), whereas phycocyanin 645 (PC645) was extracted from Chroomonas mesostigmatica (CCMP269). The watersoluble phycobiliproteins were extracted through freezing and thawing (−20 and 4 °C) in the dark. Further purification steps required centrifugation to remove any impurities following successive ammonium sulfate precipitation (40, 55, and 80%). The final centrifugation at 80% was performed in an ultracentrifuge for 20 min at 35 000 rpm, and the resulting protein pellet was resuspended in 0.050 M phosphate buffer (pH 7.5) for storage at −20 °C. Prior to experiments being run,

Ip

Ibalanced =

Sp

−

Iu Su

Iu Su

(2)

where Ip and Iu are the intensities of the pumped and unpumped probe signal, respectively, and Sp and Su are the corresponding intensities of the photodiode signals. The values entered into this equation are the sum of the four-pulse sequence and averaged for 300 consecutive cycles. This process was repeated for each delay time between −0.4 and 4 ps and repeated for five trials.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpclett.6b02237. 4728


Letter


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Frequency domain data filtration procedure, protein steady-state spectra (absorption and fluorescence), and Fourier transform maps (PDF)

(15) Joo, T.; Jia, Y.; Yu, J. Y.; Lang, M. J.; Fleming, G. R. Third-Order Nonlinear Time Domain Probes of Solvation Dynamics. J. Chem. Phys. 1996, 104, 6089−6108. (16) Lang, M. J.; Jordanides, X. J.; Song, X.; Fleming, G. R. Aqueous Solvation Dynamics Studied by Photon Echo Spectroscopy. J. Chem. Phys. 1999, 110, 5884−5892. (17) Oh, M. H.; Salvador, M. R.; Wong, C. Y.; Scholes, G. D. ThreePulse Photon-Echo Peak Shift Spectroscopy and Its Application for the Study of Solvation and Nanoscale Excitons. ChemPhysChem 2011, 12, 88−100. (18) Rosenthal, S. J.; Xie, X.; Du, M.; Fleming, G. R. Femtosecond Solvation Dynamics in Acetonitrile: Observation of the Inertial Contribution to the Solvent Response. J. Chem. Phys. 1991, 95, 4715−4718. (19) Jimenez, R.; Fleming, G. R.; Kumar, P. V.; Maroncelli, M. Femtosecond Solvation Dynamics of Water. Nature 1994, 369, 471− 473. (20) Carter, E. A.; Hynes, J. T. Solvation Dynamics for an Ion Pair in a Polar Solvent: Time-Dependent Fluorescence and Photochemical Charge Transfer. J. Chem. Phys. 1991, 94, 5961−5979. (21) Stratt, R. M. The Instantaneous Normal Modes of Liquids. Acc. Chem. Res. 1995, 28, 201−207. (22) Schwartz, B. J.; Rossky, P. J. An Exploration of the Relationship between Solvation Dynamics and Spectrally Determined Solvent Response Functions by Computer Simulation. J. Phys. Chem. 1995, 99, 2953−2958. (23) Muiño, P. L.; Callis, P. R. Hybrid Simulations of Solvation Effects on Electronic Spectra: Indoles in Water. J. Chem. Phys. 1994, 100, 4093−4109. (24) Nishiyama, K.; Yamaguchi, T.; Hirata, F.; Okada, T. Polar Solvation Dynamics: A Combination of the Reference Interaction-Site Model and Mode-Coupling Theories. Pure Appl. Chem. 2004, 76, 71. (25) Eom, I.; Joo, T. Polar Solvation Dynamics of Coumarin 153 by Ultrafast Time-Resolved Fluorescence. J. Chem. Phys. 2009, 131, 244507. (26) Fleming, G. R.; Passino, S. A.; Nagasawa, Y. The Interaction of Solutes with Their Environments. Philos. Trans. R. Soc., A 1998, 356, 389−404. (27) Joo, T.; Jia, Y.; Yu, J.-Y.; Jonas, D. M.; Fleming, G. R. Dynamics in Isolated Bacterial Light Harvesting Antenna (Lh2) of Rhodobacter Sphaeroides at Room Temperature. J. Phys. Chem. 1996, 100, 2399− 2409. (28) Passino, S. A.; Nagasawa, Y.; Joo, T.; Fleming, G. R. Three-Pulse Echo Peak Shift Studies of Polar Solvation Dynamics. J. Phys. Chem. A 1997, 101, 725−731. (29) Nagasawa, Y.; Yu, J.-Y.; Fleming, G. R. Solute−Solvent Interaction Dynamics Studied by Photon Echo Spectroscopies in Polymer Glasses. J. Chem. Phys. 1998, 109, 6175−6183. (30) Larsen, D. S.; Ohta, K.; Fleming, G. R. Three Pulse Photon Echo Studies of Nondipolar Solvation: Comparison with a Viscoelastic Model. J. Chem. Phys. 1999, 111, 8970−8979. (31) Everitt, K. F.; Geva, E.; Skinner, J. L. Determining the Solvation Correlation Function from Three-Pulse Photon Echoes in Liquids. J. Chem. Phys. 2001, 114, 1326−1335. (32) Piryatinski, A.; Lawrence, C. P.; Skinner, J. L. Vibrational Spectroscopy of Hod in Liquid D2o. V. Infrared Three-Pulse Photon Echoes. J. Chem. Phys. 2003, 118, 9672−9679. (33) Yang, M. Effect of Finite Pulse Duration in Three Pulse Photon Echo Experiments: Numerical Comparison of 3peps and S3pe. Chem. Phys. Lett. 2009, 467, 304−308. (34) Brazard, J.; Bizimana, L. A.; Gellen, T.; Carbery, W. P.; Turner, D. B. Experimental Detection of Branching at a Conical Intersection in a Highly Fluorescent Molecule. J. Phys. Chem. Lett. 2016, 7, 14−9. (35) Rafiq, S.; Dean, J. C.; Scholes, G. D. Observing Vibrational Wavepackets During an Ultrafast Electron Transfer Reaction. J. Phys. Chem. A 2015, 119, 11837−11846. (36) Rafiq, S.; Scholes, G. D. Slow Intramolecular Vibrational Relaxation Leads to Long-Lived Excited-State Wavepackets. J. Phys. Chem. A 2016, 120, 6792−6799.

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Gregory D. Scholes: 0000-0003-3336-7960 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS C.C.J. is supported by the Natural Science & Engineering Research Council. This work was supported by the Natural Sciences and Engineering Research Council of Canada and the United States Air Force Office of Scientific Research (FA955013-1-0005). P.A.K. and J.A.C. were supported in part by National Science Foundation Grant CHE-1213406.

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REFERENCES

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