Ultrafast Spectroscopy of Protein Dynamics - Journal of Chemical

Waters Symposium: Lasers in Chemistry

Ultrafast Spectroscopy of Protein Dynamics R. M. Hochstrasser Department of Chemistry, University of Pennsylvania, 3301 Spruce Street, Philadelphia, PA 19104-6323

Background Ten years after the publication of the first picosecondtime-scale visible spectra of the photolysis of liganded heme proteins (1), ultrafast infrared spectroscopic methods began to be applied to the same biological questions. Now transient infrared methods provide a bond-specific approach to the determination of key features of biological structures with femtosecond time resolution. The development of transient IR as applied to biology has mushroomed since 1987, when the first experiments (2), although already quite primitive by today’s standards, demonstrated the dissociation of the carbon monoxide from carbonmonoxy myoglobin through changes in the IR absorption of the carbonyl with picosecond time resolution. These measurements were soon followed by femtosecond-time-scale studies of the loss of CO by hemoglobin (3), through which it was concluded that the CO is captured on subpicosecond time scales at protein sites within a few angstroms from the heme iron. In these experiments the CO ligand was observed through its infrared spectrum both before and after its photodissociation from the iron. During this period the methods of employing infrared pulses to estimate the dynamics of structural features such as bond angles were first described (2, 4 ). The most recent and accurate results are the measurements of the angle between the CO bond and the heme plane in hemoglobin (5 ) and myoglobin (6 ), which are both close to 90°; the latter result is in sharp contrast to the much smaller angle obtained from X-ray diffraction. Another early result of transient IR spectroscopy concerned direct measurement of the flipping of the CO from an iron to a copper site after optical excitation (7 ). In the past few years the scope of applications has increased to include studies of the vibrational responses of both the cofactors and the protein in bacteriorhodopsin (8, 9) and the reaction centers from photosynthetic bacteria (10–12), and further and more revealing examinations of the CO ligand dynamics of myoglobin (13, 14). The use of transient IR to detect energy flow out of proteins into water (15) provided new information about protein dynamics and it was this experiment that led us to the notion of using IR probes to measure the transient temperature of water. When this knowledge was combined with the subsequent development of picosecond-temperature-jump methods, the conformational changes of proteins were proposed to be measurable through the changes in the amide-I infrared absorption spectra (16, 17 ) and a fruitful new technique for the study of macromolecular dynamics (17, 18) was put into place. We also will see that transient IR has proved useful not only for the study of vibrational states but also for the discovery of new electronic transitions in the infrared such as in the examples of the reaction centers dynamics (19) and bacteriorhodopsin (20). The field of transient infrared spectroscopy of biological systems has been discussed in a number of recent articles (21–24). While much of the work cited above required femtosecond pulses to expose some of the behavior, our applications

of transient spectroscopy to biology are not confined to “femtobiology”. Answers to questions of importance in biology will require the concatenation of experimental results recorded over many orders of magnitude of time. In that regard an important feature is that a single experimental configuration might now be used to examine systems evolving from the earliest moments (the femto-regime) where the stage is prepared by a light pulse, through the evolution of complexity toward time scales that are highly relevant to biological function (such as the microregimes or milliregimes). The experiments described in this article utilize ultrashort pulse lasers in both the infrared and optical regions and over a wide range of time delays, as required. Ultrafast processes of interest in chemistry and biology have traditionally been studied by means of transient visible or ultraviolet (UV) spectroscopic methods. In the condensed phases at ambient temperatures electronic spectra tend to be rather diffuse because of fast electronic dephasing processes. Nevertheless, the vibrational structure of the measured broad bands often can be deduced from the coherent dynamical response to impulsive Raman-like excitations. However, the information obtained is restricted to modes that are excited during the electronic transition and hence have Franck– Condon factors. A different set of conditions exists for transient vibrational spectroscopy. The characteristic times obtained from the widths of vibrational transitions are in the range 100 fs to a few picoseconds. Thus spectral resolution suitable for structural analysis is obtainable on these time scales. Even if the system were changing faster than this, the structural knowledge could be deduced from the observed time dependence. Transient infrared (IR) spectroscopy is complimentary to optical methods because vibrational information is obtained on modes that are not necessarily directly involved with electronic transitions. Thus a wide range of new phenomena can be addressed by this approach. These include ground-state nuclear motion in solutions and liquids, chemical reactions GaSe GaSe Terahertz TerahertzMethods Methods

AgGaS 2 AgGaS 2 KTP KTP BBO BBO

-1 Frequency Frequency(cm (cm )-1 )

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100 100

1000 1000 Fundamental Fundamental vibrations vibrations

"Translations" "Translations" Librations Librations

10000 10000

Overtone Overtone vibrations vibrations

Figure 1. Current laser techniques allow us to obtain femtosecond infrared pulses over a broad range of frequencies by using optical parametric amplifiers. Different nonlinear materials lead to pulses in different regions of the spectrum, as shown above. Pulses in the terahertz region of the spectrum can be generated by pumping semiconductor materials.

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including the characterization of vibrationally excited product states, surface reactions, and the responses of components of proteins and nucleic acids to functional processes. The potential for significant new knowledge is great in this area of research because of the existing robust spectrum–structure relationships that have been established by conventional IR spectroscopy. Almost any part of the IR spectrum can now be made available for ultrashort pulse experiments by means of nonlinear optical methods, as indicated in Figure 1. Although a wide variety of infrared experiments can now be accomplished on complex biological systems, there remains the very important set of problems related to finding theoretical descriptions of the vibrational modes of biological assemblies such as peptides (25), proteins, and nucleic acids (26 ). This article describes some recent applications of ultrafast laser techniques to biological systems, with emphasis on the use of infrared pulses. The descriptions simply highlight the main features of the results and their significance. We first describe recent experiments (19) aimed at obtaining information about the electronic transitions of the reaction center that lie in the infrared. Continuing with photosynthetic systems, we turn to a description of our studies of the exciton in the dimer of the light-harvesting complex, LH-1 (27), and on the dynamics of bacteriorhodopsin (20) as observed with nearinfrared pulses. Finally, recent examples of optical triggering of conformational changes in proteins are discussed. In these cases it is not a chromophore that is used for the initiation but a temperature jump (17) or a photochemical trigger (28).

10 9 8

22

Ground State Spectrum

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Excited State (P* ) Spectrum

Electronic Transitions of the Reaction Center during Electron Transfer as Seen in the Infrared Spectrum (19 ) The bacterial photosynthetic reaction center (RC) of Rhodobacter sphaeroides is a membrane-bound protein that contains eight chromophores approximately arranged around a noncrystallographic C2 axis (29– 31): two bacteriochlorophyll-a molecules (PL and PM) forming the so-called special pair, two accessory bacteriochlorophyll-a molecules (BL and BM), two bacteriopheophytin-a molecules (HL and HM), and two quinones (QM and QL). In a molecular dimer AB the monomeric excited state is split into states that are symmetric (+) and antisymmetric (–) linear combinations of A*B and AB*. The splitting is twice the magnitude of the coupling between A*B and AB*. These | + 冔 and | – 冔 states are often called “exciton states”. The lowest excited singlet state of the RC is the lowest component of a pair of excitonic states, PY᎑ and PY+, that are formed from the two singlet QY transitions on the two bacteriochlorophyll-a molecules that make up the special pair. On the basis of experimental evidence (32, 33) and electronic structure calculations (34 –38), the lowest energy component of the pair of excitonic levels is PY᎑. It is the spectrum of PY᎑ — or P*, as it is often called—that lives for only 3 ps and is shown in Figure 2 along with the groundstate spectrum of the reaction center. Considerable uncertainty remains, however, regarding the higher-lying states of the RC. Excitation of the special pair in the lowest absorption band of the RC at ~870 nm leads to a series of fast electron transfers with very high quantum efficiency down the L branch of the RC but not down the M branch (39 – 48). This indicates that the effective symmetry of the electron-transferring system is lowered from C2. To understand this fast and unidirectional electron transfer in the RC, it is crucial to understand the nature of the electronic states involved in this process.

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Figure 2. The excited-state spectrum of the photosynthetic reaction center (left), obtained by femtosecond infrared spectroscopy, is compared here with the ground-state absorption spectrum (right).The two bands in the infrared spectrum are not present in the groundstate spectrum. Putative positions of electronic states of the reaction center are depicted in the center of the diagram, where BL, BM, HL, and HM are the two accessory bacteriochlorophylls and two bacteriopheophytins, PY ᎑, PY+, PX ᎑, PX+ are the Y- and X-derived excitonic states of the special pair, and CT refers to states with charge transfer character. The scale is in units of 1000 cm᎑1.

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P + 2720 cm-1 Y PY -

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+

- +

(P H )

2600 cm-1 3.4 ps 900 nm

(P + H-)-

Figure 3. Diagram of the excited states of the photosynthetic reaction center at different stages after optical excitation, as inferred from transient infrared spectroscopy. In our experiments, a femtosecond pulse at 810 nm excites the accessory bacteriochlorophylls, which transfer the energy to the special pair in 120 fs. Electron transfer follows and takes ~3 ps. Low-lying electronic states of the intermediates are probed with infrared femtosecond pulses. B* and B** are the first and second excited states of the accessory bacteriochlorophylls, PY ᎑ and PY+ are the lower and upper excitonic states of the special pair, and P+H᎑ depicts the charge separated reaction center, where a hole transfer excitation can occur in the special pair.

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Dynamical Properties of the Dimer from the Light Harvesting Complex ( LH1) The antenna systems of photosynthetic organisms are responsible for harvesting solar energy over a broad spectral region and transferring the excitation to RCs to initiate charge separation (49). Photosynthetic purple bacteria contain two types of antenna proteins; the core (LH-1) complex, which surrounds each RC, and peripheral (LH-2) complexes, which are interspersed between them. Recent structural data (50, 51) have revealed that both complexes possess symmetric ringlike structures that can be described as circular oligomers of a basic repeating subunit. In the present study, we explored the internal dynamics of the subunit, using femtosecond pump-probe spectroscopy to investigate the isotropic and anisotropic transient kinetics of the B820 dimer from LH-1 complexes of Rhodospirillum rubrum G9. The primary goal

of this work is to identify and follow processes within the subunit that will influence the mechanism of energy migration in antenna aggregates. Excitation of B820 with 800-nm, 35-fs pulses is expected to prepare a coherent superposition of the dimer | + 冔 and | – 冔 eigenstates (in a ratio of approximately 2:1 obtained from the convolution of the pumping light pulse with the dimer spectrum). Dephasing of this superposition state will yield a distribution of populations in the two levels, which will subsequently equilibrate to establish the Boltzmann distribution (90% in the | + 冔 state at room temperature). The observed anisotropy is shown in Figure 4 along with a structure of the LH-1 dimer. The anisotropy in a transient absorption experiment (21) is the difference between the absorption for probe pulses polarized parallel and perpendicular to the pumping pulse, normalized to the isotropic absorption. The anisotropy measures changes in the orientation of the electronic transition dipoles. In the present case the dipoles undergo a damped oscillation of their direction in the laboratory space as the excitation moves back and forth between the two halves of the dimer. A summary of the conclusions from these experiments is contained in the cartoon of Figure 5, which describes the energy levels of the dimer and various relaxation rates that are consistent with the observations made. The electronic dephasing is faster than 20 fs, and the dimer exciton dephasing is about 40 fs. A near strongly damped oscillation of 65 fs shows up in the measurement, as a result of there being a coherent superposition of the | + 冔 and | – 冔 states created by the light pulse. The populations of the dimer levels equilibrate in 500 fs. The isotropic signal shows oscillations due to periodic vibrational motion. Such effects are similar to those seen in intact light-harvesting complexes (18, 20, 52–54). These findings imply that although the initially prepared state of the dimer is delocalized over the whole dimer, the dimer nuclear motion can initiate significant localization of the excitation energy in the light-harvesting complex, which consists of an assembly of dimers.

Figure 4. Left: structure of the dimeric subunit from the peripheral (LH-2) antenna complex of Rhodopseudomonas acidophila (McDermott et al. Nature 1995, 374, 517–521) showing the orientation of B850 bacteriochlorophyll pigments bound to transmembrane helical polypeptides. Right: pump-probe anisotropy of the bacteriochlorophyll dimer within the B820 subunit of core (LH-1) antenna complexes from Rhodospirillum rubrum, measured using 35-fs excitation and probe pulses (centered at 800 nm).

Figure 5. Schematic depiction of the electronic and vibrational coherences which are created and probed during the femtosecond pump-probe measurements on the B820 dimer subunit. Excitation with 800 nm, 35 fs pulses creates superpositions of the |+冔 and |᎑ 冔 dimer exciton states, as well as of the Franck–Condon active vibrations within each electronic level; Γex and Γvib represent the rates of electronic and vibrational dephasing, respectively, and τeq is time constant for equilibration between the exciton levels.

The bands observed in the transient absorption spectrum of P* are new and have no equivalent in the ground state absorption spectrum of the RC, and their interpretation is not yet established with certainty. The broad band centered at 5300 cm᎑1 most likely consists of more than one transition. The excited states clearly have charge resonance character, in which the configurations PL+ PM᎑ and PL᎑ PM+ are mixed into the exciton wave functions PL*PM and PLPM*. The 2710 cm᎑1 band also has a charge transfer source for its dipole strength. The 1900 cm᎑1 band is suggested to be related to a | PY᎑ 冔 to |PY+ 冔 transition. A summary of the kinetics and the excited electronic states that have been observed in our experiments is shown in Figure 3. The results present sharp challenges for theoretical electronic structure calculations, which are of course needed for a full interpretation of the experimental results. Infrared spectroscopy was also used to examine the changes in the vibrational spectrum of RCs during the electron transfer (11, 12). One of the more interesting observations (11) was of the changes in the amide carbonyl bands from the electric fields (Stark effects) arising from charge separation.


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Near Infrared Probing of Bacteriorhodopsin Suggests a Revision of the Potential Surface for the Initial Nuclear Motions Bacteriorhodopsin (bR) is a retinal protein that acts as a light-driven proton pump in the cell membrane of halobacteria (55). The photocycle of bR is one of the most studied biological photoinitiated reactions (56 ). The primary process in the cycle is the all-trans to 13-cis isomerization of the retinal moiety of bR. The kinetics of formation of the first few intermediates in the cycle (57–60) are usually depicted in the following scheme: hv

500 fs

3ps

BR570 → BR* → J625 → K590

the common picture of the photochemical process in this molecule. According to this picture, impulsive excitation by means of a vertical transition forms a wave packet on the excited state potential surface (see Fig. 6), which undergoes a 200-fs nuclear motion that would be observable as a timedependent shift in the emission spectrum. However, our new results indicate there is a very fast (< 50 fs) initial motion but no slow motions as demanded by the conventional potential surface. Instead an intermediate state with a lifetime of ≈ 700 fs is formed. This configuration is considered to be the transition state of the isomerization reaction. Optical Triggering of Protein Conformational Change

The subscripts in this scheme refer to the maximum absorption wavelength in nanometers for each species (61). BR* is the first excited state of the all-trans retinal in bR. It is thought that the J intermediate is already an isomerized retinal (62), while K is the product of further vibrational and conformational relaxation. The detailed form of the reactive potential and the dynamics of the motion of the isomerizing retinal on it are not known yet, although resonance Raman experiments (63–67) and visible ultrafast experiments (59, 60, 68–73) have provided important contributions towards the construction of the model potential surface shown in Figure 6. By probing the bR photoisomerization reaction in the near infrared we were able to identify a new excited state absorption band, centered at ≈ 750 nm (20). The occurrence of this band leads to cancellation of the contribution of stimulated emission to the transient spectra, and it should therefore carry a rather large oscillator strength. The time dependence of the shape of the stimulated emission or transient absorption signals of bR (see Fig. 7) suggests the need to reconsider

Questions concerning the physical and chemical nature of protein folding are among the most challenging in biological research (74 –77). Internal motions of macromolecules such as rotations about single bonds, chemical exchange reactions, diffusion over molecular dimensions, and barrier-crossing processes can occur on nanosecond or even picosecond time scales, so protein structure reorganization is expected to involve ultrafast intermediate steps. Some of the faster processes in protein folding might involve relatively small alterations in overall structure. Therefore the probes used to examine them must be sensitive to subtle changes in, for example, nonbonded interactions, alterations in the weaker chemical bonds, charge distributions, and motions of pieces of the structure. For this reason we decided to use an infrared (IR) spectroscopic method (21, 23, 24), which is sufficiently structure sensitive at a chemical-bond resolution, to identify any ultrafast steps (17) in conformational dynamics. The IR spectra of proteins in the region of the amide vibrations of the polypeptide structures are well known to be sensitive to the state of the protein. For example, there

Figure 6. Traditional potential energy surfaces for the isomerization reaction of bacteriorhodpsin, showing a putative 200-fs motion on the excited state surface, followed by a 500-fs crossing back to the ground state accompanied by the cis-trans isomerization. One prediction from these surfaces is that the wavelength of fluorescence from the excited state surface will shift with time, as depicted by the broken arrows. Our measurements show that this does not happen, pointing to a need for a revision in the common picture of the reaction dynamics in this molecule.

Figure 7. Pump-probe signals after excitation of bacteriorhodopsin (bR) at 400 nm. At this wavelength, excess vibrational energy leads to nonexponential reaction dynamics. The kinetic scheme for the isomerization reaction of bR is shown, with lifetimes derived from the measurements. Absorption from the first product state ( J ) is detected at 640 nm; stimulated emission from the excited bR, which decays as the reaction proceeds, is seen at 900 nm. Excited-state transient absorption is detected at 754 nm.

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are distinct differences between the IR spectra of random coil, α-helical, β-sheet, β′-sheet, and turn structures of polypeptides (25). These differences arise from the dependence of interactions between the various amide groups on the local polypeptide structures. One can therefore conceive of carrying out time-resolved IR capable of following the kinetics of structure change as it affects these different spatial regions of the polypeptide backbone. Experiments on the kinetics of folding also require that the system be triggered to suddenly change. For this purpose we developed an ultrafast temperature jump (T-jump) method (16, 17). In another approach (28) we investigated optical triggering of protein conformational changes. In one example the early events of folding were studied by photoinitiation of α-helix formation in de novo peptides in which a disulfide bond links the ends of a 17-amino-acid polypeptide chain, thereby constraining it to a more randomly coiled form. The absorption of a UV pulse caused femtosecond time-scale dissociation of the –S–S– bond. The recombination of the thiyl radicals measured for delay times t from 1 ps to 10 µs is approximately described by a stretched exponential with an exponent of 0.1, so that the instantaneous rate of recombination decays approximately with 1/t over the whole seven orders of magnitude in time. These recombination kinetics may be a consequence of the continuous scaling of the diffusional dynamics by the formation of the α-helix. At the longest times the helix is presumed to be formed (helix formation occurs on the ca. 100-ns time scale according to recent measurement [18]), but its ends still can undergo very slow recombination as a result of the structure fluctuations. Ultrafast infrared spectroscopy again proved useful for this study of helix dynamics. The amide carbonyls were seen to undergo shifts during photodissociation of the S–S bond. We have concluded that the shift is caused by the sudden change in electric field at the carbonyl when the radical is formed and the sulfur atoms acquire a partial charge. Summary and Prospects The importance of physicochemical methods to the unraveling of problems in biology is easily documented and many research groups are involved in this field. However, biological systems are represented by structural and electronic motifs that are frequently not easy to mimic in chemical systems. The conditions can be sufficiently unique in biology that novel ideas, principles, and interpretative methods will probably need to be developed to describe them. Therefore, biological questions, besides being important to address in their own right, present a still relatively untapped fertile source of exciting opportunities for contributions to the field of chemical physics. Ultrafast phenomena in biology are a typical example. The field was initiated in the 1970s when ultrafast was limited by technology to picosecond, and since then many remarkable and exciting discoveries have been made. This progress was certainly influenced by advances in laser technology leading to new kinds of ultrafast spectroscopy, such as the infrared methods discussed here. Throughout this period strong connections were made between the responses of biological systems and the ongoing advances in light-induced condensed-phase dynamics involving chemical reactions— particularly isomerization and atom-transfer reactions, energy relaxation, and coherent phenomena. As shorter pulse experi-

ments became possible, new phenomena began to be observed in experiments on condensed-phase small molecules and biological systems. The fundamental issues in both areas concern the profound influences of the environment on the processes of interest, whether they be electron transfers, energy transfers, or chemical reactions. In the future, as the time resolution available for experiments becomes still shorter and the equilibrium molecular and environmental nuclear motions are frozen for a more significant portion of the measurement, we might begin to see nonadiabatic solvent dynamics (where the coupled solvent motions and reactive dynamics are on similar time scales), electronic effects, and signatures of nuclei subject to very large forces undergoing irreversible processes on very steep potential energy surfaces. It is interesting to conjecture whether strong connections could then be made with the results of comparable experiments on isolated molecule systems undergoing reactions, should they become available, since these remarkably fast steps are likely to be determined by the internal forces alone. However, even in this regime the observed dynamics of biological systems will depend on the response of a distribution of forces arising from the intrinsic inhomogeneity of the condensed-phase system at sufficiently short times. The final solutions to these questions must come from a combination of advances in theoretical and experimental methods involving studies over a broad range of frequencies and times. In this presentation, I summarize a few recent experimental measurements made by my research group. The wide range of topics was chosen to illustrate the extraordinary utility of the transient spectroscopy measurement—in particular, but not exclusively, those with infrared femtosecond pulses—to many different questions in biology. Each of these subjects is a small part of a major research field involving other research groups, references to which can be more fully obtained from the cited papers. Acknowledgments This research was supported by NIH and NSF. The research discussed here was carried out with my research group (S. Gnanakaran, E. Gooding, G. Haran, Y. Kholodenko, R. Kumble, Y. Mizutani, S. Palese, M. Volk, and K. Wynne); collaborators (M. Chance, W. F. DeGrado, P. L. Dutton, H. Lu, C. Moser, R. Visschers, and A. Xie); and University of Pennsylvania/NIH Research Resource staff (M. Phillips). I wish to express my gratitude to them all. Literature Cited 1. Greene, B. I.; Hochstrasser, R. M.; Weisman, R. B.; Eaton, W. A. Proc. Natl. Acad. Sci. USA 1978, 75, 5525. 2. Moore, J. N.; Hansen, P. A.; Hochstrasser, R. M. Chem. Phys. Lett. 1987, 138, 110. 3. Anfinrud, P. A.; Han, C.; Hochstrasser, R. M. Proc. Natl. Acad. Sci. USA 1989, 86, 8387–8391. 4. Hansen, P. A.; Moore, J. N.; Hochstrasser, R. M. Chem. Phys. 1989, 131, 49. 5. Locke, B.; Lian, T.; Hochstrasser, R. M. Chem. Phys. 1991, 158, 409. Locke, B.; Lian, T.; Hochstrasser, R. M. Chem. Phys. 1995, 190, 155. 6. Lim, M.; Jackson, T. A.; Anfinrud, P. A. Science 1995, 269, 962. 7. Dyer, R. B.; Einarsdottir, O.; Killough, P. M.; Lopez-Gariga, J. J.; Woodruff, W. H. J. Am Chem. Soc. 1989, 111, 7657.


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Waters Symposium: Lasers in Chemistry 8. Diller, R.; Cowen, B. R.; Maiti, S.; Iannone, M.; Bogomolni, R.; Hochstrasser, R. M. Biophys. J. 1992, 61, A533. 9. Diller, R.; Maiti, S.; Walker, G. C.; Cowen, B. R.; Pippenger, R.; Bogomolni, R. A.; Hochstrasser, R. M. Chem. Phys. Lett. 1995, 241, 109. 10. Maiti, S.; Cowen, B. R.; Diller, R.; Iannone, M.; Moser, C.; Dutton, P. L.; Hochstrasser, R. M. Biophys. J. 1992, 61, A100. 11. Maiti, S.; Walker, G. C.; Cowen, B. R.; Pippenger, R.; Moser, C. C.; Dutton, P. L.; Hochstrasser, R. M. Proc. Natl. Acad. Sci. USA 1994, 91, 10360-10364. 12. Hamm, P.; Zurek, M.; Mäntele, W.; Scheer, H.; Zinth, W. Proc. Natl. Acad. Sci. USA 1995, 92, 1826-1830. 13. Lim, M.; Jackson, T. A.; Anfinrud, P. A. J. Chem. Phys. 1995, 102, 4355. 14. Lim, M.; Jackson, T. A.; Anfinrud, P. A. J. Phys. Chem. 1996, 102, 12043. 15. Lian, T. S.; Locke, R. B.; Kholodenko, Y.; Hochstrasser, R. M. J. Phys. Chem. 1995, 98, 11648. 16. Phillips, C. M.; Mizutani, Y.; Hochstrasser, R. M. Biophys. J. 1994, 66A, 398. 17. Phillips, C. M.; Mizutani, Y.; Hochstrasser, R. M. Proc. Natl. Acad. Sci. USA 1995, 92, 7292. 18. Williams, S.; Causgrove, T. P.; Gilmanshin, R.; Fang, K. S.; Callender, R. H.; Woodruff, W. H.; Dyer, R. B. Biochemistry 1996, 35, 691-697. 19. Wynne, K.; Haran, G.; Reid, G. D.; Moser, C. C.; Dutton, P. L.; Hochstrasser, R. M. J. Phys. Chem. 1996, 100, 5140–5148. 20. Haran, G.; Wynne, K.; Xie, A.; He, Q.; Chance, M.; Hochstrasser, R. M. Chem. Phys. Lett. 1996, 261, 389–395. 21. Hochstrasser, R. M. Ultrafast Vibrational Spectroscopy of Molecular and Protein Dynamics; in Monographs on Chemistry in the 21st Century Series, IUPAC; El-Sayed, M. A., Ed.; Blackwell: Boston, 1995. 22. Cowen, B. R.; Hochstrasser, R. M. In Bioanalytical Application of FTIR Spectroscopy; Mantsch, H. H.; Chapman, D., Eds.; Wiley: New York, 1995; pp 107–129. 23. Walker, G. C.; Hochstrasser, R. M. In Laser Techniques in Chemistry; Meyers, A. B.; Rizzo, T. R., Eds.; Wiley: New York, 1995; pp 385–422. 24. Locke, B.; Diller, R.; Hochstrasser, R. M. In Biomolecular Spectroscopy; Clark, R. J. H.; Hester, R. E., Eds.; Wiley: New York, 1993; pp 1– 47. 25. Krimm, S.; Bandekar, J. Adv. Protein Chem. 1986, 38, 181–364. 26. Infrared Spectroscopy of Biomolecules; Mantsch, H.; Chapman, D., Eds.; Wiley: New York, 1996. 27. Kumble, R.; Palese, S.; Visschers, R. W.; Dutton, P. L.; Hochstrasser, R. M. Chem. Phys. Lett. 1996, 261, 396–404. 28. Volk, M.; Kholodenko, Y.; Gooding, E.; Lu, H.; DeGrado, W. F.; Hochstrasser, R. M. J. Phys. Chem. B 1997, 101, 8607–8616. 29. Deisenhofer, J.; Epp, O.; Miki, K.; Huber, R.; Michel, H. J. Mol. Biol. 1984, 180, 385–398. 30. Deisenhofer, J.; Epp, O.; Miki, K.; Huber, R.; Michel, H. Nature 1985, 318, 618. 31. El-Kabbani, O.; Chang, C. H.; Tiede, D.; Norris, J.; Schiffer, M. Biochemistry 1991, 30, 5361. 32. Breton, J. Biochem. Biophys. Acta 1985, 810, 235–245. 33. Zinth, W.; Knapp, E. W.; Fischer, S. F.; Kaiser, W.; Deisenhofer, J.; Michel, H. Chem. Phys. Lett. 1985, 119, 1. 34. Interpretation of Optical Reaction Center Spectra; Scherer, P. O. J.; Fischer, S. F., Eds.;CRC: Boca Raton, FL, 1991. 35. Parson, W. W.; Warshel, A. J. Am. Chem. Soc. 1987, 109, 6152. 36. Thompson, M. A.; Zerner, M. C.; Fajer, J. J. Phys. Chem. 1991, 95, 5693. 37. Thompson, M. A.; Zerner, M. C. J. Am. Chem. Soc. 1991, 113, 8210. 38. Won, Y.; Friesner, R. A. J. Phys. Chem. 1988, 92, 2208. 39. Wraight, C. A.; Clayton, R. K. Biochem. Biophys. Acta 1973, 333, 246. 40. Woodbury, N. W.; Becker, M.; Middendorf, D.; Parson, W. W. Biochemistry 1985, 24, 7516. 41. Martin, J. L.; Breton, J.; Hoff, A. J.; Migus, A.; Antonetti, A. Proc. Natl. Acad. Sci. USA 1986, 83, 957–961.

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42. Breton, J.; Martin, J.-L.; Migus, A.; Antonetti, A.; Orszag, A. Proc. Natl. Acad. Sci. USA 1986, 83, 5121–5125. 43. Breton, J.; Martin, J.-L.; Fleming, G. R.; Lambry, J.-C. Biochemistry 1988, 27, 8276–8284. 44. Kirmaier, C.; Holten, D. Proc. Natl. Acad. Sci. USA 1990, 87, 3552–3556. 45. Chan, C. K.; DiMagno, T. J.; Chen, L. X. Q.; Norris, J. R.; Fleming, G. R. Proc. Natl. Acad. Sci. USA 1991, 88, 11202. 46. Schmidt, C. A.; Kaiser, W.; Lauterwasser, C.; Meyer, M.; Scheer, H.; Zinth, W. Proc. Natl. Acad. Sci. USA 1990, 90, 11757. 47. Vos, M. H.; Jones, M. R.; Hunter, C. N.; Breton, J.; Lambry, J.C.; Martin, J.-L. Biochemistry 1994, 33, 6750–6757. 48. Schmidt, S.; Arlt, T.; Hamm, P.; Huber, H.; Naegele, T.; Wachtweitl, J.; Meyer, M.; Scheer, H.; Zinth, W. Chem. Phys. Lett. 1994, 223, 116. 49. van Grondelle, R.; Dekker, J. P.; Gillbro, T.; Sundstrom, V. Biochem. Biophys. Acta 1994, 1187, 1. 50. Karrasch, S.; Bullough, P. A.; Ghosh, R. EMBO J. 1995, 14, 631. 51. McDermott, G.; Prince, S. M.; Freer, A. A.; HawthornthwaiteLawless, A. M.; Papiz, M. Z.; Cogdell, R. J.; Isaacs, N. W. Nature 1995, 374, 517. 52. Visser, H. M.; Somsen, O. J. G.; van Mourik, F.; Lin, S.; van Stokkum, I. H. M.; van Grondelle, R. Biophys. J. 1995, 69, 1083. 53. Bradforth, S. E.; Jimenez, R.; van Mourik, F.; van Grondelle, R.; Fleming, G. R. J. Phys. Chem. 1995, 99, 16179. 54. Jimenez, R.; Dikshit, S. N.; Bradforth, S. E.; Fleming, G. R. J. Phys. Chem. 1996, 100, 6825 (1996). 55. Oesterhelt, D.; Stoeckenius, W. Proc. Natl. Acad. Sci. USA 1973, 70, 2853. 56. Mathies, R. A.; Lin, S. W.; Ames, J. B.; Pollard, W. T. Annu. Rev. Biophys. Biophys. Chem. 1991, 20, 491–518. 57. Polland, H.-J.; Franz, M. A.; Zinth, W.; Kaiser, W.; Koelling, E.; Oesterhelt, D. Biophys. J. 1986, 49, 651–662. 58. Petrich, J. W.; Breton, J.; Martin, J. L.; Antonetti, A. Chem. Phys. Lett. 1987, 137, 369–375. 59. Dobler, J.; Zinth, W.; Kaiser, W.; Oesterhelt, D. Chem. Phys. Lett. 1988, 144, 215. 60. Pollard, W. T.; Cruz, C. H. B.-Ë.; Shank, C. V.; Mathies, R. A. J. Chem. Phys. 1989, 90, 199. 61. Lozier, R. H.; Bogomolni, R. A.; Stoeckenius, W. Biophys. J. 1975, 15, 955. 62. Doig, S. J.; Reid, P. J.; Mathies, R. A. J. Phys. Chem. 1991, 95, 6372–6379. 63. Myers, A. B.; Harris, R. A.; Mathies, R. A. J. Chem. Phys. 1983, 79, 603. 64. Atkinson, G. H.; Brack, T. L.; Blanchard, D.; Rumbles, G. Chem. Phys. 1989, 131, 1. 65. van den Berg, R.; Jang, D. L.; Bitting, H. C.; El-Sayed, M. A. Biophys. J. 1990, 58, 135. 66. Ujj, L.; Bolodin, B. L.; Popp, A.; Delaney, J. K.; Atkinson, G. H. Chem. Phys. 1994, 182, 291. 67. Shreve, A. P.; Mathies, R. A. J. Phys. Chem. 1995, 99, 7285. 68. Blanchard, D.; Gilmore, D. A.; Brack, T. L.; Lemaire, H.; Hughes, D.; Atkinson, G. H. Chem. Phys. 1991, 154, 155. 69. Du, M.; Fleming, G. R. Biophys. Chem. 1993, 48, 101–111. 70. Kandori, H.; Yoshihara, K.; Tomioka, H.; Sasabe, H.; Shicida, Y. Chem. Phys. Lett. 1993, 211, 385. 71. Logunov, S. L.; Song, L.; El-Sayed, M. A. J. Phys. Chem. 1994, 98, 10674. 72. Birge, R. R.; Findsen, L. A.; Pierce, B. M. J. Am. Chem. Soc. 1987, 109, 5041. 73. Robb, M. A.; Bernardi, F.; Olivucci, M. Pure Appl. Chem. 1995, 67, 783. 74. Englander, S. W.; Mayne, L. Annu. Rev. Biophys. Biomol. Struct. 1992, 21, 243. 75. Kim, P. S.; Baldwin, R. L. Annu. Rev. Biochem. 1990, 59, 631–660. 76. Sali, A.; Shakhnovich, E.; Karplus, M. Nature 1994, 369, 248–251. 77. Skolnick, J.; Kolinski, A.; Godzik, A. Proc. Natl. Acad. Sci. USA 1993, 90, 2099–2100.


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