Unusual Temperature Dependence of Photosynthetic Electron

Dec 30, 2008 - Analysis of the Kinetics of PHA Recombination in Membrane-Embedded Wild-Type and Mutant Rhodobacter sphaeroides Reaction Centers betwee...
2 downloads 8 Views 1MB Size
Download PDF
818

J. Phys. Chem. B 2009, 113, 818–824

Unusual Temperature Dependence of Photosynthetic Electron Transfer due to Protein Dynamics Haiyu Wang,†,‡ Su Lin,†,‡ Evaldas Katilius,†,‡ Christa Laser,†,‡ James P. Allen,‡ JoAnn C. Williams,‡ and Neal W. Woodbury*,†,‡ The Biodesign Institute at Arizona State UniVersity, 1001 South McAllister AVenue, Tempe, Arizona 85287-5201, and The Department of Chemistry and Biochemistry, Arizona State UniVersity, Tempe, Arizona 85287-1604 ReceiVed: August 20, 2008; ReVised Manuscript ReceiVed: October 16, 2008

The initial electron transfer rate and protein dynamics in wild type and five mutant reaction centers from Rhodobacter sphaeroides have been studied as a function of temperature (10-295 K). Detailed kinetic measurements of initial electron transfer in Rhodobacter sphaeroides reaction centers can be quantitatively described by a reaction diffusion formalism at all temperatures from 10 to 295 K. In this model, the time course of electron transfer is determined by the ability of the protein to interconvert between conformations until one is found where the activation energy for electron transfer is near zero. In reaction centers with a free energy for electron transfer similar to wild type, the reaction proceeds at least as fast at cryogenic temperatures as at room temperature. This may be because interconversion between conformations at low temperature is restricted to conformations with near zero activation energy (it is not possible to diffuse away from this region of conformational space). In contrast, mutants with a decreased free energy initially find themselves in conformations unfavorable for electron transfer and require more extensive conformational diffusion to achieve a low activation energy conformation. They therefore undergo electron transfer more slowly at 10 K vs 295 K. Introduction Photosynthetic reaction centers convert light into chemical energy by a multistep electron transfer process, and represent one of the most facile systems for the study of protein-mediated electron transfer in biology. In reaction centers of Rhodobacter sphaeroides, the initial energy conserving reaction involves picosecond electron transfer from an excited electron donor, P* (P is a pair of bacteriochlorophylls), via a neighboring bacteriochlorophyll, BA, to a bacteriopheophytin, HA (Figure 1).1 Previously, we have suggested that the electron transfer kinetics for this reaction is dictated by conformational diffusion of the protein, rather than by a static activation barrier,2 an idea that builds on previous molecular dynamics simulations modeling of electron transfer in reaction centers3,4 and other proteinmediated reactions.5,6 This description of protein-mediated reactions is embodied mathematically in the reaction diffusion model, using a generalized Smoluchowski equation.6-8 A similar approach has previously been used to describe reactions such as CO binding to heme proteins,6 barrierless isomerization reactions,7 solvent effects on electron transfer reactions,8 and enzyme catalysis.9 In order to apply this to electron transfer reactions, the reorganization energy is divided into two parts, λf and λp. λf is the reorganization energy associated with solvent/protein motion that takes place faster than electron transfer, such that all possible conformations associated with this motion are effectively explored before electron transfer occurs (corresponding to the * Corresponding author. E-mail: [email protected]. Phone: 480-9653294. Fax: 480-727-0396. † The Biodesign Institute at Arizona State University. ‡ The Department of Chemistry and Biochemistry, Arizona State University.

traditional solvent reorganization energy in the Marcus formalism10). λp is the energy associated with reorganization that involves protein motion on a time scale that is equal to or slower than electron transfer and is therefore unable to explore all of the conformational space before electron transfer occurs. When all of the key conformational states can be explored effectively instantaneously relative to the reaction, the reaction is sometimes described as ergodic, while reactions that involve conformational space that cannot be entirely explored on time scales faster than electron transfer are kinetically controlled by conformational diffusion and can be considered nonergodic.11 Thus, dividing the reorganization energy into λf and λp is one way of treating nonergodic reactions, by formalizing the dichotomy of time scales associated with the conformational changes important to electron transfer and treating those motions that are rapid relative to electron transfer differently from those that are not. The key mechanistic concept behind the reaction diffusion model is that once the excited state is created by light absorption, conformational diffusion proceeds (in some sense, the effective reorganization energy varies) until the free energy and the effective reorganization energy are nearly equal and opposite; at this point, the activation energy given by the Marcus term, (∆G° + λ)2/4λ, goes to zero (∆G° is the free energy difference, and λ is the Marcus theory reorganization energy10), and electron transfer rapidly takes place. This mechanism lends a robust quality to photosynthetic electron transfer; even if environmental conditions result in changes in the energetics of the system, the local environment varies dynamically until the activation energy is minimal. Previous measurements of the P* decay kinetics (initial electron transfer kinetics) coupled with measurements of tryptophan absorbance changes (protein conformational diffusion) in a series of reaction center mutants at room temperature with

10.1021/jp807468c CCC: $40.75  2009 American Chemical Society Published on Web 12/30/2008

Photosynthetic Electron Transfer

Figure 1. Cofactor arrangement of wild type reaction centers from Rhodobacter sphaeroides. P is a bacteriochlorophyll dimer. BA and BB are bacteriochlorophyll monomers. HA and HB are bacteriopheophytins. QA and QB are ubiquinones. The amino acid residues that were altered to vary the free energy for electron transfer are shown in shades of blue, violet, green, yellow, and orange. The mutant L168HE introduces a potentially charged residue in the vicinity of P; L131LH and M160LH alter hydrogen bonding to P; L153HS changes the original histidine that coordinates the Mg in BA, making it harder to reduce; M203GL has altered a glycine between P and BA that is thought to affect the state of a bound water molecule in that region; and M210YF apparently varies the energetics of both the electron donor and acceptor (refs 9 and 21-24).

a wide range of ∆G° demonstrated that it is possible to quantitatively describe the complex dynamics of all of these samples using the reaction diffusion model, varying only the driving force between mutants.2 Furthermore, the free energies obtained in this way for the different mutants are in agreement with past measurements.2,12 Here, the ability of this model to describe the unusual temperature dependence of electron transfer as a function of free energy is explored. One of the most surprising aspects of the initial electron transfer reaction in photosynthesis is that it continues unabated as the temperature is lowered down to liquid helium temperatures. In fact, both the first and second electron transfer steps in the reaction center from Rhodobacter sphaeroides accelerate slightly as the temperature is dropped.13 Most of the previous explanations for the temperature dependence of initial electron transfer have involved either an activation energy that was static and near zero14-17 or, more recently, electron transfer from a system in which vibrational relaxation is incomplete.18,19 However, the reaction diffusion model described above gives a different picture of what controls the electron transfer rate. In this mechanism, protein conformational dynamics explores the energy landscape, and this process dictates the rate of electron transfer, even at cryogenic temperatures. This raises some important issues. Electron transfer in photosynthetic organisms has been taking place for billions of years. One tends to think of this process as being highly optimized for rate and yield by evolution, and at some levels, such as placement of cofactors and the general electrostatic environment provided by the protein, this is clearly the case. However, rapid electron transfer at 4 K is clearly not an evolutionary constraint for photosynthesis and there is no reason to think that an energy landscape optimized for physiological temperature would remain optimal

J. Phys. Chem. B, Vol. 113, No. 3, 2009 819 at liquid helium temperatures. It is certainly the case that artificial electron transfer systems that work efficiently at both room temperature and cryogenic temperatures have proven difficult to construct.20,21 For most solvents, the decrease in the reorganization energy as the temperature is lowered is large, and the ability of the solvent to reorient around the charge separated state and thus stabilize it is greatly limited. Therefore, the balance between reorganization energy and free energy is shifted as the temperature is lowered, resulting in a barrier to electron transfer at cryogenic temperatures. This implies that the efficiency of photosynthetic electron transfer at cryogenic temperatures must be inherent in the mechanism of photosynthetic electron transfer, rather than something specifically optimized by evolution. Does the ability to mediate electron transfer at low temperature reflect an intrinsic property of protein as a heterogeneous, dynamic solvent, or is this a very specific property of the reaction center? Are there, in fact, appropriate protein dynamics available in the reaction center to facilitate electron transfer at cryogenic temperatures, or does some other mechanism dominate at low temperature? In order to address these questions, it was necessary to both measure the dynamics of protein conformational diffusion and carefully measure the kinetics of initial electron transfer as a function of temperature between 10 and 295 K. Materials and Methods Mutants. The construction and expression of all of the mutants, L168H f E (L168HE), L131L f H + M160L f H (L131LH + M160LH), L153H f S (L153HS), M210Y f F (M210YF), and M203G f L (M203GL), were described previously (refs 12 and 22-25). Quinones were removed using published methods.26 The resulting reaction centers were suspended in 15 mM Tris-HCl (pH 8.0), 0.025% LDAO, and 1 mM EDTA and then mixed with 67% (v/v) glycerol. The samples were placed between two glass plates with a rubber spacer of 1.2 mm and then attached to the coldfinger of a closed circulated helium displex (APD) that can control the temperature between 300 and 10 K. In our previous room temperature study (ref 2), 15 mutants were studied. For low temperature measurements, the samples had to have the quinones removed to avoid buildup of long-lived charge separated states in the stationary samples. Many of the mutants used at room temperature were difficult to prepare in this way. Thus, five of the most stable mutant RCS that covered a broad range of electron transfer times (refs 12 and 22-25) were selected for this study. In the case of the double mutant, L131LH + M160LH, which was important to measure because of its long electron transfer time, it was only possible to measure the P* decay using only the limited amount of available sample after quinone removal but not the 280 nm signal, which is a much more sample- and timeconsuming measurement. Transient Spectroscopy. Femtosecond transient absorption spectroscopy following the stimulated emission of P* was performed as follows. A titanium sapphire oscillator (Tsunami, Spectra-Physics) was used to generate 100 fs, 800 nm laser pulses at a repetition rate of 82 MHz. These pulses were used to seed an optical amplifier system (Spitfire, Spectra-Physics), resulting in pulses of approximately 0.9 mJ at a repetition rate of 1 kHz. Part of the pulse energy (∼10%) was then used to generate a white light continuum by focusing the beam into a sapphire plate, and this was used to create the sample and reference probe beams. The remainder of the amplified 800 nm pulse was used to pump an optical parametric amplifier (OPA800, Spectra-Physics) generating excitation pulses at a wave-

820 J. Phys. Chem. B, Vol. 113, No. 3, 2009

Wang et al.

length of 860 nm (second harmonic of idler beam). Transient absorption changes were measured at 930 nm (stimulated emission of P* state) using a monochromator (SP150, Action Res. Corp.) and a diode detector (model 2032, New Focus Inc.). The relative polarization of the excitation and probe beams was set to the magic angle. For 800 nm pump and 280 nm probe measurements of the Trp absorbance changes, the pump and probe beams were reversed, and 800 nm was used as the excitation. The 280 nm probe beam was obtained by doubling 560 nm generated by the OPA using a BBO crystal. The relative polarization of the excitation and probe beams was parallel. The effective system excitation pulse width was about 150 fs. Data Analysis. The model used to analyze the electron transfer dynamics was based on the reaction diffusion equation.8 In this model, the reactant and product energy surfaces involve both fast, q, and slow, x, reaction coordinates associated with protein motion. The potentials are given by

1 Vreact ) q2 + Vp(x) 2 1 Vproduct ) (q - qproduct)2 + Vp(x - xproduct) + ∆G° 2

(1)

where Vp(x) is the free energy as a function of position along the reaction coordinate, x (we take this to be a harmonic potential, Vp(x) ) 0.5x2). The total reorganization energy λ is 2 the sum of λf ) 0.5qproduct (fast nuclear modes) and λp ) 2 0.5xproduct (slow protein contribution). Assuming that the fast nuclear modes are effectively instantaneous on the time scale of electron transfer, the reaction diffusion equation as a function of the slow protein coordinate is given by8,27,28

[

]

dVp(x) ∂F(x, t) ∂ ∂ k T + F(x, t) - K(x) F(x, t) ) Dp(t) ∂t ∂x B ∂x dx (∆G° + λ - 2x√λp)2 J2 π where K(x) ) exp p λfkBT 4λfkBT (2)



[

]

where F(x,t) is the population distribution in the reactant state, J is the electronic coupling matrix element, ∆G° is the free energy gap between the reactants and the products, and kB and T are the Boltzmann constant and temperature, respectively. Dp(t) is the time-dependent diffusion coefficient that can be obtained from the protein dielectric relaxation function Cp(t), Dp(t) ) -Cp(t)-1 dCp(t)/dt.27,28 Cp(t) can be empirically determined by monitoring the tryptophan absorbance change during the electron transfer.2 Finally, the observable time-dependent population of the electron donor, P*(t), is given by

P*(t) )

∫ F(x, t) dx

(3)

For the purposes of this analysis, the above equations can be rearranged into a more convenient form:

Let

X)

x

√kBT

- X0,

where

(

) [



[ ]

X0 ≡

∆G° + λ

√4λpkBT

; thus,

dCp(t) ∂ ∂ ∂F(X, t) ) -Cp(t)-1 + 2(X + X0) × ∂t dt ∂X ∂X F(X, t) - K(X)F(X, t) λp J2 π K(X) ) exp - X2 p λfkBT λf P*(t) )

∫ F(X, t) dX

]

(4)

To fit the temperature-dependent data, we use the semiclassical Hopfield formula to modify eq 4, involving a single vibrational mode.14,29 For this, the terms 2λfkBT and 2λpkBT in eq 4 are replaced with λfpω coth(pω/2kBT) and λppω coth(pω/2kBT), respectively. A simple equilibrium distribution is assumed for the initial condition in which F(X, 0) ∼ exp[-(X + X0)2].30 The temperature-dependent protein relaxation function Cp(t) is obtained by fitting the composite tryptophan decay kinetics at each temperature. The fits of the electron transfer kinetics (stimulated emission decays) are obtained by performing a numerical solution of eq 4, adjusting the parameters ω, X0, J, λp, and λf. A global data analysis was carried out using locally written software developed under a MATLAB environment (Mathworks Inc.). Results and Discussion Temperature-Dependent Protein Dynamics. As described in detail in ref 2, we measured protein conformational dynamics by monitoring the tryptophan absorbance changes at 280 nm after excitation of the reaction center cofactors using 800 nm light (Figure 2). The kinetic traces in Figure 2 were normalized such that the initial and final values of each decay are set the same, allowing comparison of the decay kinetics over the intervening time range. The shape of the curve changed with temperature as described below (Figure 2A) but was essentially invariant from mutant to mutant (Figure 2B), as had been observed previously at room temperature.2 The absolute amplitude of the signal was more difficult to evaluate between samples and temperatures, though it generally became somewhat smaller with decreasing temperature for all samples, consistent with a freezing out of certain collective protein motions and an associated decrease in reorganization energy. As the temperature was lowered from 295 to 250 K, there were only minor changes in the tryptophan absorbance kinetics on the picosecond time scale. However, below 250 K (the approximate glass transition under these conditions), a substantial change occurred in the kinetic profile. One can see that the relative amplitude of the protein motion on time scales longer than about 30 ps (presumably the collective motion of the protein matrix) is greatly reduced relative to the motion observed on time scales faster than about 30 ps (local harmonic motion). In contrast, the faster conformational diffusion continues even at 10 K. We also measured the tryptophan absorbance changes upon 860 nm excitation for wild type; it gave essentially the same results (data not shown). Temperature-Dependent Electron Transfer. Detailed measurements of the electron transfer kinetics were performed in the same set of reaction center samples as a function of temperature by monitoring the decay of the stimulated emission signal from P* at 930 nm (Figures 3 and 4). For wild type and for the two mutants L168HE and L153HS, the overall electron transfer dynamics accelerates somewhat at lower temperature. L168HE has a free energy for electron transfer greater than wild

Photosynthetic Electron Transfer

Figure 2. (A) Temperature-dependent protein dynamics determined by monitoring the changes in the tryptophan absorbance band at 280 nm in quinone-removed wild type reaction centers. (inset) The same traces shown on a shorter time scale. (B) Comparison of the transient absorbance change in the tryptophan absorbance band at 280 nm in quinone-removed reaction centers from wild type and four mutants at 10 K. (inset) The same traces shown on a shorter time scale. The mutants are described in the legend to Figure 1.

Figure 3. Temperature-dependent electron transfer kinetics for wild type and five reaction center mutants determined from the transient absorbance changes at 930 nm (stimulated emission from P*) using 860 nm excitation. The mutants are described in the legend to Figure 1. The solid red lines are fitting results using the reaction diffusion model. For all samples, the kinetics at 50 K is nearly identical with that at 10 K.

type,22 and L153HS has a slightly smaller free energy.23 In contrast, the mutants M210YF, M203GL, and L131LH + M160LH, which all have much smaller free energy values than wild type,12,24,25 show an overall slower decay of the stimulated emission at cryogenic temperatures. In addition, the electron transfer kinetics of these three mutants become more distinctly heterogeneous compared to room temperature, maintaining some fast (picosecond) kinetic components but picking up very slowly decaying components as well, that stretch out to hundreds of picoseconds or longer. This bifurcated behavior of the mutants, with the fastest, highest free energy mutants speeding up with decreasing temperature and the slower mutants decreasing further in rate, correlates with the observed temperature-

J. Phys. Chem. B, Vol. 113, No. 3, 2009 821 dependent dynamics of conformational diffusion as monitored by the tryptophan absorbance changes (Figure 2). Mutants with picosecond decay times can take advantage of the protein motion that is still abundant on that time scale at low temperature (evidenced by the large Trp absorbance change signal on the 0-30 ps time scale in Figure 2). However, mutants for which picosecond conformational diffusion times are insufficient must utilize the more restricted protein motion on longer time scales at cryogenic temperature. Reaction Diffusion Model. To explore the ability of the reaction diffusion model to quantitatively describe the temperature dependence of the electron transfer kinetics, the data in Figures 3 and 4 were fit to a modified model, involving a single vibrational mode, as described in Materials and Methods. The reaction diffusion model describes the electron transfer kinetics as a function of the free energy, λf, λp, a coupling term, and the vibrational frequency. First, to get an overall picture, the fitting parameters determined previously for room temperature were used as initial guesses (∆G° for wild type was initially set to 200 meV).2 The temperature-dependent kinetics was then fitted individually, varying all parameters. Very little change was observed in ∆G° as a function of temperature for wild type and each of the mutants (about 20 meV). In contrast, the total reorganization energy for each mutant decreases substantially with decreasing temperature. On the other hand, comparing different mutants at a particular temperature, there was relatively little change in the total reorganization energy (about 30 meV). Therefore, it became apparent that, at least approximately, the free energy changed between mutants (as expected) but the reorganization energy was nearly constant and the reorganization energy changed with temperature but the free energy for each mutant remained roughly constant. In these initial fits where all parameters were free, the electronic coupling matrix elements varied over a range between 25 and 40 cm-1, but there was no clear correlation with mutant or temperature. The fitting always gave a vibrational mode frequency of pω ) 165 cm-1, which is close to the main vibrational mode of 150 cm-1 observed via coherence spectroscopy.31 Given these results, a second fitting was performed. In this fit, the reaction free energy was allowed to vary from mutant to mutant, but for any given mutant, it was held essentially independent of temperature (∆G was allowed to vary over a very small range with temperature, normally a few percent; see Table 1). In contrast, the values of two reorganization energy terms, λf and λp, were allowed to vary as a function of temperature but were kept constant at any particular temperature between the different mutants (Table 1). All other parameters in the fit were held constant both between mutants and as a function of temperature. Clearly, such fits could have been performed by allowing more parameters to vary (in particular, the reorganization terms and the free energy can play against each other to some extent), but this more constrained fit was sufficient to obtain a nearly quantitative description of the electron transfer kinetics of all of the mutants as a function of temperature (Figures 3 and 4, solid lines, parameters given in Table 1). This is remarkable given the almost opposite temperature-dependent trends of the kinetics for the fast and slow mutants. There have been a number of other treatments of the temperature dependence of photosynthetic electron transfer,15,32-35 including some that consider specifically the molecular dynamics of the reaction center structure.17,19,36 However, until now, it has not been possible to correlate detailed measurements of electron transfer kinetics with measurements of protein dynamics, an approach that has proven to be critical for creating a

822 J. Phys. Chem. B, Vol. 113, No. 3, 2009

Wang et al.

Figure 4. Temperature-dependent electron transfer kinetics of the slower mutants M210YF, M203GL, and L131LH + M160LH shown on a several hundred picosecond time scale. The solid red lines are fitting results using the reaction diffusion model.

TABLE 1: Free Energy Associated with the Initial Electron Transfer Reaction for Various Mutants and Temperature-Dependent Reorganization Energies as Determined via Fitting with a Reaction Diffusion Model (J ) 39 cm-1 and pω ) 165 cm-1) λf (meV), λp (meV) sample

∆G° (meV)

10 K

90 K

150 K

200 K

250 K

295 K

L168HE WT L153HS M210YF M203GL L131LH + M160LH

-233 ( 15 -197 ( 3 -150 ( 3 -103 ( 5 -97 ( 3 -64 ( 5

120, 120

126, 120

155, 120

179, 115

210, 104

280, 70

complete description of the temperature and free energy dependence of photosynthetic electron transfer. We should point out that such a fit is not unique. It was also possible to accurately fit the data using significantly smaller absolute values of the free energy and the total reorganization energy. However, if this was done, ∆G° became quite temperature dependent, especially for the slow mutants, decreasing by about 40 meV as the temperature was lowered. Figure 5 shows the temperature dependence of the reorganization energy parameters that resulted from the fits described above. As the temperature decreases from 295 to 100 K, λf drops by more than a factor of 2. In contrast, λp actually increases

Figure 5. Temperature dependence of λf (circles), λp (squares), and λtotal (triangles) reorganization energies resulting from fitting with a reaction diffusion model for the wild type and five mutants (the reorganization energy values were held the same for all of the mutants at each temperature in the fit).

slightly down to 200 K and then remains approximately constant at lower temperatures. The increase in λp with decreasing temperature presumably arises as protein dynamics that is rapid compared to the rate of electron transfer at room temperature and slows down into the regime where it is comparable to the electron transfer rate (no longer able to explore the available conformational space completely on time scales faster than electron transfer). The observation that substantial picosecond time scale conformational motion remains at low temperature, as evidenced by the early time tryptophan absorbance changes of Figure 2B, is consistent with the notion that, as the temperature decreases, the large-scale, collective motions of proteins are diminished while small-scale, local protein motions are much less affected.37 Apparently, the local, rapid movements that still take place at cryogenic temperature explore conformational space sufficiently to allow electron transfer to take place on the picosecond time scale in most mutants. Relationship between Free Energy Landscape and Protein Function. As demonstrated above, the reaction diffusion formalism, coupled with measurements of the dynamics of conformational diffusion in the reaction center protein, provides a surprisingly robust, and conceptually simple, mechanistic description of the detailed kinetics of photosynthetic electron transfer. For the wild type and the five mutants studied, a complete picture of the temperature-dependent dynamics is provided by varying only the free energy (as a function of the mutation) and the components of the reorganization energy (as a function of temperature). This model suggests that one can

Photosynthetic Electron Transfer think of photosynthetic electron transfer in terms of an energy landscape associated with the process of conformational diffusion during the reaction. Reaction centers, such as wild type, that start near the region of the conformational space in which the free energy and the effective reorganization energy balance, can rapidly diffuse to the point where electron transfer can take place with little or no activation energy. As long as there are pathways through this local region of conformational space that do not require crossing large barriers, the diffusion will take place rapidly at low temperature as well as at room temperature. In fact, decreasing the temperature can give rise to an acceleration of the reaction, as is observed, by limiting conformational diffusion along unproductive pathways. On the other hand, reaction center mutants that start far from the region of conformational space where the activation energy is low (for example, due to a large change in free energy for the reaction) will demonstrate slower and more complex electron transfer dynamics at low temperature. This arises because barriers to conformational diffusion that were not significant at room temperature become insurmountable and must be traversed. These concepts are completely consistent with the fact that the high free energy mutants continue to have fast electron transfer rates (and even speed up) at low temperature while the low free energy mutants became slower. Very recently, LeBard et al.36 have performed extensive molecular dynamics simulations of Rhodobacter sphaeroides reaction centers, effectively using this simulation instead of the Trp absorbance signal as a way of exploring the conformational changes in the reaction center. From this, they were able to directly calculate the reorganization energy parameters and use these results in the reaction diffusion formalism to accurately simulate our previous data at room temperature. In addition, they were able to predict the overall temperature dependence of the electron transfer rate over a limited temperature range.36 This merger of structurally based dynamics simulations with the reaction diffusion formalism suggests that a complete description, that explicitly includes the dynamics of the reaction center protein structure, should be possible for the wild type and mutant reaction center electron transfer reactions as a function of temperature. It is hard to imagine that there was any significant evolutionary pressure to create a system that would effectively traverse the low temperature energy landscape. Instead, the robust temperature independence of the electron transfer reaction appears to reflect an inherent property of the reaction center protein and perhaps of other protein environments: proteins behave like heterogeneously dynamic dielectric materials, with the agility to explore conformational space under many different conditions, searching for a minimum in the activation energy of a reaction. In support of this, there is recent evidence that, in reaction center mutants that undergo electron transfer to the B-side cofactors (P* to P+HB-), the rate of this alternate electron transfer reaction also increases by roughly a factor of 2 with decreasing temperature.38 As far as is currently known, this reaction pathway is not used in the native system and therefore presumably not optimized by evolution, again suggesting that the dynamic properties of the protein that facilitate rapid electron transfer are not specific to a particular, highly evolved, structural region. Interestingly, Gray’s laboratory has reported that electron transfer in a Ru-azurin complex increases in rate slightly at low temperatures compared to room temperature,39 suggesting that dynamics may play a role in the temperature dependence of other electron transfer proteins as well. Molecular dynamics simulations as well as certain dynamics measurements of

J. Phys. Chem. B, Vol. 113, No. 3, 2009 823 enzymatic reactions also indicate that the ability of proteins to dynamically mediate reaction energetics may not be unique to electron transfer or to picosecond time scales.40,41 The photosynthetic reaction center simply provides a facile experimental system in which conformational diffusion and reaction kinetics can be directly compared. Acknowledgment. This work was supported by NSF grants MCB0642260 and MCB064002. The laser equipment used in this work was purchased with funds from NSF grant BIR9512970. The authors would like to thank Steven Boxer for the gift of M210YF strain and Dmitry Matyushov for helpful discussions and access to submitted work. References and Notes (1) Woodbury, N. W.; Allen, J. P. In Anoxygenic Photosynthetic Bacteria; Blankenship, R. E.; Madigan, M. T.; Bauer, C. E., Eds.; Kluwer Academic Publishers: Dordrecht, The Netherlands, 1995; Vol. 2, pp 527557. (2) Wang, H.; Lin, S.; Allen, J. P.; Williams, J. C.; Blankert, S.; Laser, C.; Woodbury, N. W. Science 2007, 316, 747–750. (3) Warshel, A.; Chu, Z. T.; Parson, W. W. Science 1989, 246, 112– 116. (4) Gehlen, J. N.; Marchi, M.; Chandler, D. Science 1994, 263, 499– 502. (5) Austin, R. H.; Beeson, K. W.; Eisenstein, L.; Frauenfelder, H.; Gunsalus, I. C. Biochemistry 1975, 14, 5355–5373. (6) Agmon, N.; Hopfield, J. J. J. Chem. Phys. 1983, 79, 2042–2053. (7) Bagchi, B.; Fleming, G. R.; Oxtoby, D. W. J. Chem. Phys. 1983, 78, 7375–7385. (8) Sumi, H.; Marcus, R. A. J. Chem. Phys. 1986, 84, 4894–4914. (9) Min, W.; Xie, X. S.; Bagchi, B. J. Phys. Chem. B 2008, 112, 454– 466. (10) Marcus, R. A.; Sutin, N. Biochim. Biophys. Acta 1985, 811, 265– 322. (11) Matyushov, D. V. Acc. Chem. Res. 2007, 40, 291–301. (12) Lin, X.; Murchison, H. A.; Nagarajan, V.; Parson, W. W.; Allen, J. P.; Williams, J. C. Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 10265– 10269. (13) Huber, H.; Meyer, M.; Scheer, H.; Zinth, W.; Wachtveitl, J. Photosynth. Res. 1998, 55, 153–162. (14) Hopfield, J. J. Proc. Natl. Acad. Sci. U.S.A. 1974, 71, 3640– 3644. (15) Bixon, M.; Jortner, J. AdV. Chem. Phys. 1999, 106, 35–202. (16) Warshel, A.; Parson, W. W. Q. ReV. Biophys. 2001, 34, 563–679. (17) Parson, W. W.; Warshel, A. J. Phys. Chem. B 2004, 108, 10474– 10483. (18) Haffa, A. L. M.; Lin, S.; Katilius, E.; Williams, J. C.; Taguchi, A. K. W.; Allen, J. P.; Woodbury, N. W. J. Phys. Chem. B 2002, 106, 7376–7384. (19) Parson, W. W.; Warshel, A. Chem. Phys. 2004, 296, 201–216. (20) Liddell, P. A.; Kuciauskas, D.; Sumida, J. P.; Nash, B.; Nguyen, D.; Moore, A. L.; Moore, T. A.; Gust, D. J. Am. Chem. Soc. 1997, 119, 1400–1405. (21) Kuciauskas, D.; Liddell, P. A.; Lin, S.; Stone, S. G.; Moore, A. L.; Moore, T. A.; Gust, D. J. Phys. Chem. B 2000, 104, 4307–4321. (22) Williams, J. C.; Haffa, A. L. M.; McCulley, J. L.; Woodbury, N. W.; Allen, J. P. Biochemistry 2001, 40, 15403–15407. (23) Katilius, E.; Babendure, J. L.; Lin, S.; Woodbury, N. W. Photosynth. Res. 2004, 81, 165–180. (24) Treynor, T. P.; Yoshina-Ishii, C.; Boxer, S. G. J. Phys. Chem. B 2004, 108, 13523–13535. (25) Potter, J. A.; Fyfe, P. K.; Frolov, D.; Wakeham, M. C.; van Grondelle, R.; Robert, B.; Jones, M. R. J. Biol. Chem. 2005, 280, 27155– 27164. (26) Okamura, M. Y.; Isaacson, R. A.; Feher, G. Proc. Natl. Acad. Sci. U.S.A. 1975, 72, 3491–3495. (27) Okuyama, S.; Oxtoby, D. W. J. Chem. Phys. 1986, 84, 5824. (28) Hynes, J. T. J. Phys. Chem. 1986, 90, 3701–3706. (29) Hoffman, B. M.; Ratner, M. A. Inorg. Chim. Acta 1996, 243, 233– 238. (30) Okada, A. J. Phys. Chem. A 2000, 104, 7744–7750. (31) Vos, M. H.; Lambry, J.-C.; Robles, S. J.; Youvan, D. C.; Breton, J.; Martin, J.-L. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 8885. (32) Jia, Y.; DiMagno, T. J.; Chan, C.-K.; Wang, Z.; Du, M.; Hanson, D. K.; Schiffer, M.; Norris, J. R.; Fleming, G. R.; Popov, M. S. J. Phys. Chem. 1993, 97, 13180–13191.

824 J. Phys. Chem. B, Vol. 113, No. 3, 2009 (33) Huppman, P.; Arlt, T.; Penzkofer, H.; Schmidt, S.; Bibikova, M.; Dohse, B.; Oesterhelt, D.; Wachtveit, J.; Zinth, W. Biophys. J. 2002, 82, 3186–3197. (34) Matyushov, D. V. J. Chem. Phys. 2005, 122, 084507. (35) Torchala, M.; Kurzynski, M. J. Phys. Chem. B 2008, 112, 6508– 6512. (36) LeBard, D. N.; Kapko, V.; Matyushov, D. V. J. Phys. Chem. B 2008, 112, 10322–10342. (37) Frauenfelder, H.; Sligar, S. G.; Wolynes, P. G. Science 1991, 254, 1598–1603.

Wang et al. (38) Chuang, J. I.; Boxer, S. G.; Holten, D.; Kirmaier, C. J. Phys. Chem. B 2008, 112, 5487–5499. (39) Skov, L. K.; Pascher, T.; Winkler, J. R.; Gray, H. B. J. Am. Chem. Soc. 1998, 120, 1102–1103. (40) Yang, H.; Luo, G.; Karnchanaphanurach, P.; Louie, T. M.; Rech, I.; Cova, S.; Xun, L. Y.; Xie, X. S. Science 2003, 302, 262–266. (41) Henzler-Wildman, K. A.; Kern, D. Nature 2007, 450, 964–972.

JP807468C