Probing M-Branch Electron Transfer and Cofactor Environment in the

Alexej A. Zabelin, Valentina A. Shkuropatova, Vladimir A. Shuvalov, Anatoly Ya. Shkuropatov. FTIR spectroscopy of the reaction center of Chloroflexus ...
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J. Phys. Chem. B 2002, 106, 495-503

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Probing M-Branch Electron Transfer and Cofactor Environment in the Bacterial Photosynthetic Reaction Center by Addition of a Hydrogen Bond to the M-Side Bacteriopheophytin Christine Kirmaier,*,† Agnes Cua,‡ Chunyan He,† Dewey Holten,*,† and David F. Bocian*,‡ Department of Chemistry, Washington UniVersity, St. Louis, Missouri 63130-4899, and Department of Chemistry, UniVersity of California, RiVerside, California 92521-0403 ReceiVed: July 18, 2001; In Final Form: October 31, 2001

Subpicosecond time-resolved absorption and steady-state resonance Raman (RR) studies are reported for Rhodobacter capsulatus reaction centers (RCs) that incorporate an Asp in place of Val M131 near the ring V keto group of the M-side bacteriopheophytin (BPhM). The L-side C2-symmetry analogue of residue M131 is Glu L104, which is known to form a hydrogen bond to the ring V keto group of BPhL in the wild-type RC. The effects of the V(M131)D mutation were probed in the triple mutant V(M131)D/G(M201)D/L(M212)H, which also incorporates an Asp at M201 near the L-side bacteriochlorophyll (BChlL) as well as the “beta” mutation L(M212)H that results in replacement of the native BPhL with a BChl molecule (denoted β). The primary photochemistry in the triple mutant (denoted V(M131)D-DH) is similar to that reported previously for the G(M201)D/L(M212)H double mutant (denoted DH). Upon excitation, P* decays with a time constant of 15 ps via a combination of electron transfer to the L side (70%), decay to the ground state (15%), and electron transfer to the M side to form P+BPhM- (15%). The bleaching of the QX ground-state absorption band of BPhM observed upon formation of P+BPhM- is red shifted 1-2 nm from ∼527 nm in the DH RC to ∼529 nm in the V(M131)D-DH mutant, and the BPhM anion band is shifted 20 nm from 645 to 665 nm. RR experiments reveal that the ring V keto vibration of BPhM at 1705 cm-1 in the wild-type RC downshifts to 1697 cm-1 in the V(M131)D mutant. Collectively, these results indicate that the Asp introduced at M131 forms a hydrogen bond to the ring V keto group of BPhM in both ground (neutral) and anionic states of this cofactor, and further demonstrate that M-side electron transfer to form P+BPhM- occurs in both mutants. Comparison of the effects engendered by the addition of the hydrogen to BPhM with those found previously upon removal of the Glu L104 hydrogen bond to BPhL give insights into specific and global effects of the protein on the properties of these symmetry-related cofactors.

Introduction The bacterial reaction center (RC) is a membrane-bound pigment-protein complex that has a macroscopic C2 symmetric arrangement of the L and M polypeptides and the associated bacteriochlorophyll (BChl), bacteriopheophytin (BPh), and quinone (Q) cofactors (Figure 1).1 Charge separation is initiated by excitation of the dimeric BChl primary electron donor (P) to its lowest excited singlet state (P*). Utilizing BChlL in parallel mechanisms as a discreet or virtual electron carrier, an electron is transferred from P* to the L-side BPh molecule (BPhL) in ∼4 ps, followed by electron transfer to the neighboring quinone (QA) in ∼200 ps, with an overall quantum yield of ∼1 (Figure 2A).2 Photoinduced electron transfer down the M-branch to form the transient state P+BPhM- was first observed in the Rhodobacter capsulatus G(M201)D/L(M212)H mutant (denoted DH).3a The G(M201)D mutation places an Asp near ring V of BChlL, while the His introduced by the L(M212)H mutation results in replacement of BPhL with a BChl (denoted β). The effects of the BPhL f β pigment change on the charge separation/ recombination events have been studied extensively.4 The * To whom correspondence should be addressed. † Washington University. ‡ Univeristy of California.

Figure 1. Arrangement of the RC cofactors as determined by the X-ray structures of RCs from Rb. Viridis and Rb. sphaeroides.1

working model for the DH mutant is that P+BChlL- is raised significantly in free energy by Asp M201, so as to place this state well above P+β- and probably slightly above P* (Figure 2B).5 The resulting slower electron transfer to the L side allows competing electron transfer to BPhM to occur in 15% yield. Similar photochemistry is also seen in the S(L178)K/G(M201)D/ L(M212)H triple mutant (denoted KDH), which additionally

10.1021/jp012768r CCC: $22.00 © 2002 American Chemical Society Published on Web 12/20/2001

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Kirmaier et al. V(M133)D mutant that has been prepared.12 Here we have used the DH template to produce the V(M131)D-DH triple mutant in addition to also making the V(M131)D single mutant. The goals of this work were 2-fold: (1) We sought to provide independent proof that electron transfer to the M side indeed occurs, by specifically modulating the optical characteristics of BPhM and P+BPhM-. (2) We wished to compare the optical and vibrational characteristics imparted by the addition/removal of hydrogen bonds to BPhM/BPhL as a means of further probing specific and global protein-induced environmental differences between the properties of these two cofactors. Materials and Methods

Figure 2. Schematic energy level diagrams for the (a) wild-type RC and (b) DH mutant.

incorporates a Lys near BChlM.3b In the KDH RC, the yield of electron transfer to the M branch is ∼50% higher (23%), presumably due to a lower free energy of P+BChlM- and enhanced participation of this state in M-side charge separation. An even higher P+BPhM- yield of 30% is obtained in the F(L181)Y/Y(M208)F mutant in which Phe and Tyr are swapped near BChlM and BChlL.3d A 35% yield of M-side state P+φ- is obtained in the Rb. sphaeroides H(M182)L mutant in which BChlM is replaced by a BPh (φ).33b The formation of P+BPhM- was initially diagnosed by the observation of a transient absorption decrease at ∼527 nm, ascribed to bleaching of the QX ground-state absorption band of BPhM and, in subsequent work, the resolution of the accompanying anion band of BPhM- at ∼645 nm.3 For comparison, the BPhL transient QX bleaching and anion bands in state P+BPhL- (e.g, in wild-type RCs) are at 542 and 665 nm, respectively.3b,4b These spectral distinctions between BPhL and BPhM derive significantly from a known difference in hydrogen bonding of these pigments.1,6-10 The crystal structures of Rhodopseudomonas Viridis and Rhodobacter sphaeroides RCs reveal that a Glu residue at L104 is sufficiently close to the ring V keto group of BPhL to form a hydrogen bond.1 The optical and vibrational signatures of BPhL in Rb. capsulatus RCs (where Glu L104 is conserved1e) are also consistent with hydrogen bond formation to BPhL.8,9b,10 When this hydrogen bond is removed in Rb. capsulatus and Rb. sphaeroides by changing Glu L104 to a non-hydrogen-bonding amino acid such as Leu or Val, the BPhL ground-state QX and BPhL- anion bands blue shift ∼10 and 20-30 nm, respectively, and the ring V keto vibration upshifts ∼8 cm-1.4b,8-11 These shifts do not completely compensate for the differences in the spectral features of the two BPhs in the wild-type RC, indicating that the two cofactors experience environmental asymmetries in addition to hydrogen bonding in the native protein. The M-side amino acid that is C2-symmetry related to L104 is Val M131 in Rb. capsulatus and Rps. Viridis, and Thr M133 in Rb. sphaeroides.1e Thus, no possibility exists for a hydrogen bond to the ring V keto group of BPhM by the amino acid that is symmetry-related to Glu L104, nor are there other neighboring amino acids that could do so.1 The lack of a hydrogen bond to BPhM is supported by the optical differences between BPhM and BPhL noted above, and from the differences in the ring V keto vibrations of the two cofactors seen in RR spectra of wildtype RCs (∼1705 and ∼1685 cm-1, respectively).6-11 In the present study, we have made the mutation Val M131 f Asp in Rb. capsulatus in order to introduce a hydrogen bond to the ring V keto group of BPhM, similar to the Rb. sphaeroides

Mutagenesis, RC Preparation, and Sample Conditions. An aspartic acid was introduced at M131 using the “QuikChange” site-directed mutagenesis kit from Strategene. Two templates were used: the wild-type M gene and our previously constructed G(M201)D/L(M212)H double mutant (denoted DH), both in Strategene’s Bluescribe vector. The oligonucleotide primers (two complementary primers are used in the QuikChange mutagenesis strategy) changed the GTC codon for Val at M131 to GAC for Asp. The primers also introduced silent mutations in the codons of Thr133 (ACC to ACA) and of Arg134 (CGC to CGT) that resulted in creation of an Afl III site that was used for screening of candidate mutants and subsequent cloning steps. Successful mutagenesis was established by DNA sequencing utilizing Perkin-Elmer’s “Big-Dye” kit. The 935-base-pair KpnI-BamHI M-gene fragment carrying the mutation(s) was then cloned into polyHis-pU2924. PolyHis-pU2924 is a derivative of the expression vector pU2924 originally devised by Youvan and Bylina.13a It contains seven sequential His residues added at the end of the M gene immediately before the stop codon. The polyHispU2924 vector was constructed and generously provided to us by Drs. D. Hanson and P. Laible of Argonne National Laboratory.13b Further details of the mutagenesis followed as previously described.14 His-tagged RCs were isolated via a procedure modified slightly from the standard ammonium sulfate precipitation and DEAE chromatography methods to take advantage of the presence of the poly-His tag on the protein.13b,c Following LDAO solubilization of the RCs, Qiagen Ni-NTA resin was added to the raw LDAO/RC/chromatophore solution and the mixture slowly agitated on ice for 30 min. The slurry was then poured into a small column to collect the RC-bound resin. The resin was washed with buffer (10 mM Tris pH 7.8/0.05% LDAO) until the eluent had A < 0.05 between 280 and 900 nm. At this point, the RCs were eluted with buffer containing 40 mM imidazole and then dialyzed overnight to remove the imidazole. Compared to wild-type RCs, the V(M131)D-DH RCs were obtained in only low yield and were found to decompose over several months at -80 °C. The V(M131)D mutant gave higher yields and more stable RCs than the triple mutant. The V(M131)D-DH RCs used for TA experiments were solubilized in either 10 mM Tris pH 7.8/0.05% LDAO or 10 mM phosphate pH 7.8/0.05% LDAO (with no difference in observed properties). The V(M131)D RCs used for the RR experiments were solublized in 10 mM tris (pH 8)/0.01% Triton X-100/1 mM EDTA and QA was reduced by adding a slight excess of buffered sodium dithionite solution. The occupancy of the secondary quinone (QB) was found to be 98% of the decay occurred with a time constant of ∼150 ms which can be ascribed to P+QA- charge recombination. The TA and RR experiments reported here were carried out on the poly-His V(M131)D-DH and poly-His V(M131)D RCs. Our previously published work on wild-type and DH RCs was performed on the nonpoly-His proteins, purified by traditional means. As a baseline for comparisons, we have carried out extensive subpicosecond-resolved TA experiments on poly-His DH and poly-His wild-type RCs and found no significant differences in any of the spectral or kinetic data for either RC compared with those reported previously on the respective nonpoly-His proteins.13c TA Measurements. The primary electron-transfer reactions of the V(M131)D-DH mutant were investigated on flowed samples at ∼285 K. The TA spectrometer is based on an Arion pumped regeneratively amplified Ti:sapphire OPA system operated at 10 Hz. The RCs were excited with 130 fs excitation pulses at 850 or 760 nm and probed with ∼130 fs “white-light” flashes. The excitation pulses were defocused and/or attenuated such that ∼30% of the RCs were excited on a single flash. Further details of the TA apparatus, data acquisition, and data analysis methods have been described elsewhere.15 RR Measurements. The RR measurements on the V(M131)D mutant were made on optically dense (OD ∼1.0/mm at 800 nm; RC concentration ∼35 µM), snowy samples at 25 K contained in 1 mm i.d. capillary tubes. The advantages and disadvantages of using snowy versus glassy samples have been previously discussed.16 Temperature control was achieved by mounting the sample on a cold tip of a closed cycle refrigeration system (ADP Cryogenics, DE-202 Displex). The RR spectra were obtained using a red-optimized triple spectrograph and detection system that has been previously described.7 A Ti:sapphire laser (Coherent 890) pumped by an Ar ion laser (Coherent Innova 90-6) served as the excitation source. The laser powers were typically 1.5 mW. The power density on the sample was lowered by defocusing the incident beam. The resulting photon fluxes (∼100 photons s-1 RC-1) were low enough that only a few percent of the RCs exist in photogenerated transient states. Each RR data set was obtained with 4 h of signal averaging. The data acquisition time for an individual scan was dictated by the level of background fluorescence from the sample in a particular data acquisition window (vide infra). These times ranged from 30 to 60 s. Cosmic spikes in the individual scans were removed prior to co-addition of the scans. The spectral resolution was ∼2 cm-1 in all spectral regions. The spectral data were calibrated using the known frequencies of fenchone.17 The QY-excitation RR spectra observed for both the wildtype and V(M131)D RCs are superimposed on a fluorescence background that is particularly large in the region of the highfrequency ring-skeletal and carbonyl modes (1600-1760 cm-1). This background, in conjunction with the fact that the QYexcitation RR intensity enhancements of the high-frequency modes of bacteriochlorins are generally weak,16,18,19 compromise the quality of the spectra. Therefore, all the RR spectra were acquired using the shifted-excitation Raman difference spectroscopic (SERDS) technique.20,21 The application of the SERDS method to RCs has been discussed in a number of earlier publications.16,19,20,22-25 Briefly, each data set is acquired at two excitation wavelengths that differ by a small wavenumber increment (typically 10 cm-1). [The 4 h data acquisition time indicated above is for each of the two data sets required to construct a given SERDS trace.] These data sets are subtracted

J. Phys. Chem. B, Vol. 106, No. 2, 2002 497

Figure 3. Transient absorption difference spectra, acquired using 130fs, 750-nm excitation flashes, in the region of P bleaching and P* stimulated emission.

to yield a background-free RR difference (SERDS) spectrum. The RR spectra presented herein were obtained by subtracting the initial spectrum from the shifted spectrum. The spectral window is defined by the initial spectrum and corresponds to the wavenumber axis in the figures. The normal RR spectrum is then reconstructed from the SERDS data by fitting the latter to a series of derivative-shaped functions (in this case difference bands generated from Gaussian functions) of arbitrary frequency, amplitude, and width. The frequencies marked in the figures correspond to the positions of the bands used in the fits and, thus, do not necessarily correspond to the peak maxima for overlapping bands. In addition, certain bands are marked that are not clearly resolved in the spectra. These bands are indicated because their inclusion noticeably improved the quality of the fits to the SERDS data. Results TA Spectra and Kinetics. Excitation of the V(M131)D-DH RC initially produces the excited primary donor, P*. The absorption difference spectrum of this state is characterized by bleaching of the 850-nm ground-state absorption band of P and concomitant stimulated emission (870-950 nm) from P* (0.3 ps spectrum in Figure 3). A fit of the stimulated emission decay between 890 and 920 nm to a single-exponential plus a constant gives a P* lifetime of 15 ( 2 ps (data not shown), the same value as that obtained for the DH mutant.3a In wild-type RCs, the constant magnitude of P bleaching observed during the P* lifetime and to many nanoseconds after excitation reflects the formation of P+QA- in ∼100% yield (see Figure 2A). However, in the V(M131)D-DH mutant, the amplitude of P bleaching at 3 ns is reduced ∼30% compared to its initial magnitude (Figure 3). The extent of the P-bleaching decay and thus ground-state repopulation is best monitored at 830-840 nm (on the blue side of the P band), where the contribution of P* stimulated emission is minimal or absent. The P-bleaching decay in this region can be fit to two exponentials (plus a constant) with time constants of 20 ( 5 ps and 1-4 ns,26 each of which has an amplitude 15 ( 5% of the initial P bleaching. The value of the shorter component is in good agreement with the P* lifetime obtained from decay of stimulated emission, indicating that P* decay is accompanied by ∼15% return to the ground state. The slower component is assigned below, in conjunction with data in other spectral region, to quantitative charge-recombination of P+BPhM- to the ground state (QB is absent).27 Collectively, these results (and those described below) indicate that the overall photochemistry in the V(M131)D-DH mutant is basically the same as that summarized for the DH RC in Figure 2B.

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Kirmaier et al.

Figure 6. Time profile of the absorption changes between 660 and 670 nm acquired using 130-fs flashes at 850 nm (circles). The dashed line is a fit to a single-exponential plus a constant, and the solid line is a fit to the sum of two exponentials plus a constant. Data acquired before and during the flash and during the P* lifetime have been omitted (see text). Figure 4. Transient absorption difference spectra, acquired using 130fs, 850-nm excitation flashes, in the QX and anion region for the (a) V(M131)D-DH and (b) DH mutants acquired.

Figure 5. Comparison of the BPhM QX bleaching for the (a) V(M131)D-DH and (b) DH mutants acquired 43 ps after excitation as in Figure 4.

Visible-region TA difference spectra spanning 480-710 nm are shown for the V(M131)D-DH mutant in Figure 4A. Analogous data for the DH mutant are shown for comparison in Figure 4B. The spectra at 1 ps after excitation for both mutants are characteristic of state P* and include a featureless transient absorption throughout the visible region broken by bleaching of the QX band of P at ∼600 nm. The spectra for both mutants at 43 ps contain a small but distinct bleaching in the vicinity of 530 nm, which is the region ascribed to the QX ground-state absorption band of BPhM. Close inspection of this region in Figure 5 shows that the bleaching in the DH mutant is centered at ∼527 nm and in the V(M131)D-DH mutant is centered at ∼529 nm. This small shift is not resolved in the ground-state absorption spectrum, presumably due to the underlying contribution of the carotenoid absorption. Side-byside experiments (same concentration of samples studied under identical excitation conditions) showed that the magnitudes of the QX bleachings are essentially identical in the V(M131)DDH and DH mutants. Thus, the QX-bleaching data indicate that the yield of P+BPhM- in the V(M131)D-DH mutant is the same as in the DH mutant, which previously has been shown to be 15%. This value is in agreement with the 15% amplitude of

Figure 7. Spectra of the preexponential factors of the dual exponentials fits across the entire anion absorption region for the (a) V(M131)DDH and (b) DH mutants. The squares correspond to the faster (160 ps) component and the circles correspond to the slower (1-3 ns) component.

the ∼1 ns component of P-bleaching decay reflecting P+BPhMf ground state. The anion region spectrum of the DH mutant at 43 ps (Figure 4B) has a distinct peak at ∼645 nm, while the V(M131)D-DH RC has a rather featureless transient absorption between 620 and 700 nm (Figure 4A). This difference is also seen at 3 ns. The decay kinetics across the entire anion region are dual exponential for both RCs (see Figure 6), reflecting parallel electron transfer along the L side (∼70%) and along the M side (∼15%). The faster component has τ ) 160 ( 30 ps and the lifetime of the slower component is on the order of 1-4 ns.26 The two decay components and their preexponential-factor spectra (Figure 7)28 can be assigned on the basis of the detailed analysis given previously for the DH and KDH mutants.3 The 160-ps component for both the V(M131)D-DH and DH mutants has broad absorption with two apparent maxima at ∼640 and ∼690 nm (squares in Figure 7A,B) and is associated with decay

Probing the Bacterial Photosynthetic Reaction Center

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Figure 8. QY-excitation (λex ) 750, 755, 765 nm) RR spectra of the BPhs in wild-type RCs in the region of the carbonyl and high-frequency ring-skeletal modes. In each panel, the top trace is the raw SERDS data, the second trace is the fit of the SERDS data, the third trace is the SERDS residual (observed minus fit), and the bottom trace is the RR spectrum reconstructed from the SERDS data.

Figure 9. QY-excitation (λex ) 750, 755, 765 nm) RR spectra of the BPhs in V(M131)D RCs in the region of the carbonyl and high-frequency ring-skeletal modes. In each panel, the top trace is the raw SERDS data, the second trace is the fit of the SERDS data, the third trace is the SERDS residual (observed minus fit), and the bottom trace is the RR spectrum reconstructed from the SERDS data.

of the L-side intermediate, which is largely P+β- but perhaps with some mixing with P+BChlL- (Figure 2B). The longerlived component can be attributed to M-side charge recombination P+BPhM- f ground state (QB is absent), with the BPhMabsorption of P+BPhM- clearly red-shifted from ∼645 nm in the DH mutant to the 660-670 nm range for the V(M131)DDH RC. RR Spectra. The high-frequency regions (1600-1760 cm-1) of the QY-excitation (λex ) 750, 755, and 765 nm) RR spectra of Rb. capsulatus wild-type and V(M131)D RCs are shown in Figures 8 and 9, respectively. [The V(M131)D single mutant was used for the RR studies rather than the V(M131)D-DH RCs because RR scattering from the β-cofactor in the latter mutant contribute to the RR spectra obtained with λex ) 750-765 nm,

thus complicating the analysis of the data.] Although the full data acquisition spanned 1300-1760 cm-1, only the highestfrequency region is shown because it exhibits the key spectral differences among the various RCs. In both Figures 8 and 9, the top trace is the raw (unsmoothed) SERDS data; the second trace is the fit of the SERDS data; the third trace is the SERDS residual (observed minus fit); the bottom trace is the RR spectrum reconstructed from the SERDS data. The fidelity of the fits is indicated by comparison of the SERDS and residual intensities and is more than sufficient to ascertain band positions for RR features of interest to within (2 cm-1. The RR spectra shown in Figures 8 and 9 span the QY(0,0) absorption bands of both BPh cofactors. For wild-type RCs, the QY(0,0) maxima of BPhL and BPhM are at ∼760 and ∼750

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Figure 10. Comparison of the QY-excitation (λex ) 750 nm) RR spectra of the BPhs of wild-type and V(M131)D RCs in the region of the carbonyl modes. The data are the same as those shown in the left panels of Figures 8 and 9.

nm, respectively.9 However, previous RR studies have shown that selective excitation is not possible because both BPhs contribute strongly to the spectrum with excitation throughout the 750-765 nm region.7,25 In the case of the V(M131)D mutant, the QY(0,0) band of BPhM is shifted a few nanometers to the red of its position in wild-type and closer to the QY(0,0) band of BPhL (although the exact magnitude of the shift is difficult to ascertain due to the spectral widths and overlaps). This would tend to further diminish selective excitation of one BPh versus the other. Inspection of Figures 8 and 9 reveals a general similarity of the RR spectra for the V(M131)D and wild-type RCs for a variety of different excitation wavelengths. The specific assignments for the high-frequency (1400-1760 cm-1) RR bands of the two BPh cofactors in the wild-type RC7 and a variety of genetically modified RCs25 have been previously discussed in detail and will not be reiterated herein.29 Instead, we will focus on the frequencies (and not intensities) of the key features of the spectra that distinguish BPhM in the V(M131)D and wildtype RCs, and their positions relative to BPhL.30 The key objective of the RR studies of the V(M131)D RC was to ascertain whether the replacement of Val M131 with an Asp residue affords a hydrogen bond between this amino acid and the ring V C9-keto group of BPhM. The formation of such a bond would “symmetrize” the two BPh molecules inasmuch as the C9-keto group of BPhL in wild-type RCs is known to be hydrogen bonded (to the Glu L104 residue).1,8 Hydrogen bonding interactions are clearly reflected in the frequency of the stretching vibration of the C9-keto group (νC9dO). To facilitate comparison, the reconstructed spectra (obtained with λex ) 750 nm) for the wild-type and V(M131)D RCs in the region of the C9-keto modes are reproduced in Figure 10. In the case of the wild-type RC, the νC9dO modes of BPhL and BPhM are observed at 1686 and 1705 cm-1, respectively.6b,c,7,10 The νC9dO band of BPhL is overlapped with another band at ∼1681 cm-1, which is assigned to the stretching vibration of

Kirmaier et al. the C2a-acetyl carbonyl group (νC2adO) of BPhL.7,25 In the case of the V(M131)D RC, bands assignable to the νC9dO and νC2adO modes of BPhL are also observed at ∼1686 and ∼1681 cm-1, respectively, as expected because the binding site of the L-side cofactor has not been perturbed in the V(M131)D mutant (either intentionally or unintentionally). However, the 1705cm-1 band in wild-type is absent in the mutant and a new feature appears at ∼1697 cm-1, which is assigned as the νC9dO mode of BPhM in the V(M131)D RC. This spectral pattern is also reflected in the data obtained at other excitation wavelengths (Figures 8 and 9). The ∼8-cm-1 downshift of the νC9dO of BPhM in V(M131)D versus wild-type RCs is consistent with the formation of a hydrogen bond between the ring V C9-keto group and Asp M131. In particular, hydrogen bonding generally lowers the frequency of carbonyl stretching vibrations.6c,7,10 For example, the lower frequency of the νC9dO mode of BPhL versus BPhM in wild-type RCs is known to be in part due to hydrogen bonding to the C9-keto group of the former cofactor (which is absent for the latter). This point will be considered further in the Discussion. One additional observation of note on the RR spectra of V(M131)D versus wild-type RCs is the mutation-induce upshift in the 1665-cm-1 band to ∼1671 cm-1 (Figure 10). The 1665cm-1 band is due to a νC2adO mode.25 However, previous studies could not determine whether this band is a vibration of BPhM or BPhL. The assignment of the νC2adO vibrations of the cofactors is complicated by the fact that multiple bands are observed for this mode for a given cofactor due to the existence of populations of RCs having different interactions between the C2a-acetyl group and the protein (due to low torsional barriers involving the acetyl group). The fact that the 1665-cm-1 mode is affected by the introduction of Asp M131 indicates that this feature is due to the C2a-acetyl group of BPhM. The observation that altering the hydrogen bonding interactions between the C9keto group has consequences that extend beyond the site of direct interaction is consistent with previous RR studies of other genetically modified RCs.22a,25,31 These studies have also shown that the C2a-acetyl group, which lies on the opposite side of the BPh macrocycle from the C9-keto group, is perturbed by the formation/removal of hydrogen bonds to the latter group. Discussion Incorporating an Asp near BPhM at M131 complements the first mutation ever made in the bacterial RC, namely removal of Glu L104 near BPhL.8 The results reported here show that addition of Asp M131 causes formation of a hydrogen bond to the ring V keto group of BPhM, whereas removal of Glu L104 eliminates the analogous hydrogen bond to BPhL that is present in the wild-type RC.8,9b,10 In the following, we discuss the implications of the spectral changes engendered by the addition of the hydrogen bond from Asp M131 to BPhM for the observation of electron transfer to the M side of the RC and for the effects of the protein on the properties of the symmetry related BPh cofactors. M-Side Electron Transfer and Directionality. Our previous support for M-side charge separation to produce P+BPhM(occurring in parallel with L-side charge separation) in the DH and related Rb. capsulatus RCs was based on several observations, including a key bleach at ∼527 nm and a transient absorption at ∼645 nm.3 We have found here that both features are shifted in the expected manner due to the addition of a hydrogen bond from Asp M131 to the ring V keto group of BPhM. These findings reinforce our assignment of the spectral

Probing the Bacterial Photosynthetic Reaction Center features to bleaching of the QX ground-state absorption band of BPhM and absorption due to the BPhM- anion, respectively, upon formation of P+BPhM-. Our assignment of the spectral and kinetic data to P+BPhM- formation (and decay) is also supported by our results on the DH mutation in an Rb. capsulatus carotenoidless strain and in Rb. sphaeroides32 and is consistent with other observations of M-side electron transfer.33,34 Furthermore, the V(M131D)-DH mutant shows dualexponential decay kinetics in the anion region (620-720 nm) with the spectral and kinetic characteristics appropriate for decay of the L- and M-side transient intermediates derived from parallel electron transfer down the two sides of the RC. Collectively, this body of work unequivocally demonstrates that charge separation from P* to the M-side can be elicited through appropriate genetic manipulation of the cofactors and resulting effects on the free energies of the electronic states of the RC. The results reported herein also have implications for the contributors to the directionality of charge separation. The P* lifetime (∼15 ps) and P+BPhM- yields (∼15%) in the V(M131)DDH mutant are essentially the same as in the DH RC. This result parallels the finding that removal of the hydrogen bond from Glu L104 to BPhL only slightly increases the P* lifetime from that found in wild-type RCs (∼4 to ∼6 ps), with no apparent affect on the P+BPhL- yield.8 Both of these results indicate that small (50-80 meV) changes35 in the free energies of P+BPhMand P+BPhL- do not significantly alter the effective rate constants for electron transfer to the respective sides of the wildtype RC (despite likely differences in mechanisms) and do not alter the directionality of charge separation.8 Instead, directionality is controlled primarily by other factors,36 including a substantial contribution from the relative free energies of P+BChlL- and P+BChM- with respect to P*.3,33b,37 Effect of the Protein on the Properties of the Two BPh Cofactors. Previously it was shown that the νC9dO mode of BPhL upshifts ∼8 cm-1 upon removal of the hydrogen bond to Glu L104 (1685 to 1693 cm-1).10 We have found here that the νC9dO mode of BPhM downshifts by the same magnitude within experimental uncertainty (from 1705 to 1697 cm-1) when an Asp is placed at M131 and a hydrogen bond forms. Thus, there is consistency in the opposite directions of the shifts induced by the two mutations that can be correlated with the loss/gain of a hydrogen bond. This finding and the comparable magnitudes of the shifts prompt consideration of several fundamental issues regarding hydrogen bonding involving amino acids and the tetrapyrrole cofactors in the RC. Before proceeding to these more fundamental issues, it is useful to consider other implications of finding ∼8 cm-1 shifts upon addition/removal of hydrogen bonds to the BPh cofactors. The ∼8 cm-1 hydrogen-bonding-induced frequency shifts represent only ∼40% of the ∼20 cm-1 difference between the νC9dO vibrations of BPhL and BPhM in the wild-type RC. It is also noteworthy that νC9dO vibrational frequency of BPhM in the absence of the hydrogen bond (i.e., ∼1705 cm-1 in the wildtype RC) is comparable to that observed for BPh in nonhydrogen-bonding, low-dielectric solvents in vitro.6,7 From this perspective, the ∼1685 cm-1 energy of the νC9dO mode of BPhL is (anomalously) low by ∼12 cm-1 in the wild-type RC. A significant fraction (∼60%) of the difference between the νC9dO vibrational frequencies of the two BPh cofactors in the wild-type RC can be understood in terms of a higher effective dielectric constant of the environment around BPhL versus that around BPhM (e.g.,  ∼ 2 versus 9).10 [The frequency of the νC9dO vibration is lower in vitro in higher dielectric solvents.] The assessment that dielectric effects differentially perturb the

J. Phys. Chem. B, Vol. 106, No. 2, 2002 501 L- versus M-side BPhs is further supported by the results obtained herein for the V(M131)D mutant. In particular, “symmetrization” of the hydrogen bonding interactions on BPhL and BPhM is not sufficient to yield identical frequencies for the νC9dO vibrations of the two cofactorssthe frequency for BPhL is still considerably lower. The assessment from the RR data of a higher dielectric of the microenvironment of L- versus M-side BPh is in accord with conclusions from other studies. In particular, it has been proposed from optical-Stark-effect data38 that the cofactors on the L side of the RC see a higher overall protein dielectric strength than the symmetry-related M-side pigments. Similarly, it has been calculated that the electrostatic potentials (provided by many amino acids and the protein as a whole) are different on the two sides of the RC,37c in a manner commensurate with a higher L-side dielectric. These differences in potentials/ dielectrics are thought to primarily underlie the lower free energies of the L-side charge-separated states compared to those on the M side (e.g. P+BPhL- lower than P+BPhM- and P+BChlL- lower than P+BChlM-).36h,37,39 Effects of Hydrogen-Bonding on the Optical Properties of the BPh Cofactors. The presence/absence of hydrogen bonds to the ring V keto groups of the two BPh cofactors has effects on the optical spectra40 that parallel and complement the effects on the vibrational spectra. First, we note that the ring V C9 position is very close to the Y (N1-N3) axis in the bacteriochlorin macrocycle.41 Consequently, hydrogen bonding involving the ring V keto group should predominantly affect the Y-polarized (i.e., QY) optical transitions of the cofactors (rather than the QX transitions). Thus, it is not surprising to find that manipulation of these hydrogen bonds in the M131/L104 mutants causes substantial (∼20 nm) red/blue shifts in the anion bands of BPhM/BPhL in states P+BPhM-/P+BPhL- (∼645/665 nm for the nonbonded/bonded forms of both cofactors). This point follows because the bacteriochlorion anion transition(s) in the 600-700 nm region are essentially Y polarized.42 Likewise, alteration of these hydrogen bonds also affects the QY ground-state absorption transitions of the neutral forms of the BPh cofactors, although it is difficult to pinpoint the precise magnitudes of the effects due to spectral overlap, which is severe even at low temperature.9 A priori one would expect that the QX ground-state absorption bands of the BPh cofactors should not be appreciably affected by manipulation of the hydrogen bonds to the ring V keto groups. This expectation is consistent with the finding of only a e2 nm red shift in the room-temperature ground-state QX band and transient QX bleaching of BPhM due to the V(M131)D mutation. A similarly small ∼4 nm shift is found for the corresponding V(M133)D Rb. sphaeroides mutant even at 20 K.12 In retrospect, then, what is surprising is the large ∼10 nm blue shift in the QX transition of BPhL observed upon removal of the ring V keto hydrogen bond in the Glu(L104) mutants.4b,8 This shift comprises ∼75% of the 12-15-nm difference from the position of the QX band of BPhM in wild-type RCs. The anomalously large effect on the QX band of BPhL versus BPhM may also originate from the differences in dielectric environments around the two cofactors. In particular, the larger optical shifts might be anticipated in a region of higher dielectric constant. In general, the smaller optical effects of manipulation of hydrogen bonds to BPhM versus BPhL might predict that other types of mutations on the M versus L sides may also give smaller effects on properties of the respective M- versus L-side cofactors (e.g., the free energy of the charge separated states).

502 J. Phys. Chem. B, Vol. 106, No. 2, 2002 Hydrogen Bonding of Amino Acids to the Cofactors. To a first approximation, the finding of the comparable RR shifts (∼8 cm-1, but in opposing directions) in both the V(M131)D and E(L104)L mutants relative to the wild-type RC suggests that the hydrogen bonds from Asp M131 to BPhM and from Glu L104 to BPhL have roughly similar strengths. In zeroth order (and all other things being equal), one might expect that Asp would make stronger hydrogen bonds than Glu based on the differences in pka values of these amino acids. On the other hand, the carboxylic acid group of Asp may be farther from the ring V keto group of BPhM compared to a Glu due to the shorter length of the side chain of Asp. These two effects could compensate to some extent. Additionally, the difference in the dielectric/electrostatic environments of the two cofactors comes into play. These considerations may be of fundamental importance when considering the general issue of whether either Asp or Glu residues hydrogen bond to the ring V keto groups of the BPh cofactors in the RC in the first place. Based on solution pKa values, both of these residues should be ionized, and thus incapable of hydrogen bonding. Nevertheless, the fact that hydrogen bonds are observed indicates that the Asp and Glu residues near the BPh cofactors are indeed protonated. [Similarly, both Asp and Glu appear to be protonated and hydrogen bond to the ring V keto group of P.35c] The tendency to remain protonated must ultimately derive from the fact that the free energy of the hydrogen-bonded (and thus protonated) carboxylic acid group is lower that that of the ionized group in the protein environment. This consideration may well override any distinctions between the electronic/structural differences of Glu and Asp (presuming there is sufficient structural/electronic pliability in the site to allow protonation and hydrogen bonding interactions). At present, we are continuing to pursue these fundamental issues by comparing the effects of either Asp or Glu at the M131 and L104 sites near the symmetry related BPh cofactors. The ultimate goal is to elucidate factors that dictate hydrogen bonding in the RC and other pigment protein complexes. Acknowledgment. This work was supported by Grant MCB0077187 (C.K. and D.H.) from the National Science Foundation and Grant GM-39781 (D.F.B.) from the National Institute of General Medical Sciences. References and Notes (1) (a) Ermler, U.; Fritzsch, G., Buchanan, S.; Michel, H. Structure 1994, 2, 925-936. (b) Deisenhofer, J.; Epp, O.; Sinning, I.; Michel, H. J. Mol. Biol. 1995, 246, 429-457. (c) Yeates, T. O.; Komiya, H.; Chirino, A.; Rees, D. C.; Allen, J. P.; Feher, G. Proc. Natl. Acad. Sci. U.S.A. 1988, 85, 7993-7997. (d) El-Kabbani, O.; Chang, C.-H.; Tiede, D.; Norris, J.; Schiffer, M. Biochemistry 1991, 30, 5361-5369. (e) Komia, H.; Yeates, T. O.; Rees, D. C.; Allen, J. P.; Feher, G. Proc. Natl. Acad. Sci. U.S.A. 1988, 85, 9012-9016. (2) (a) Deisenhofer, J.; Norris, J. R., Eds. The Photosynthetic Reaction Center; Academic: San Diego, 1993; Vol. II. (b) Blankenship, R. E., Madigan, M. T., Bauer, C. E., Eds. Anoxygenic Photosynthetic Bacteria; Kluwer Academic Publishers: Dordrect, The Netherlands, 1995. (c) MichelBeyerle, M. E., Ed. The Reaction Center of Photosynthetic Bacteria; Springer: Berlin-Heidelberg, 1996. (3) (a) Heller, B. A.; Holten, D.; Kirmaier, C. Science 1995, 269, 940945. (b) Kirmaier, C.; Weems, D.; Holten, D. Biochemistry 1999, 38, 11516-11530. (c) Roberts, J. A.; Holten, D.; Kirmaier, C. J. Phys. Chem. B 2001, 105, 5575-5584. (d) Kirmaier, C.; He, C.; Holten, D. Biochemistry 2001, 40, 12132-12139. (4) (a) Kirmaier, C.; Gaul, D.; DeBey, R.; Holten, D.; Schenck, C. C. Science 1991, 251, 922-927. (b) Kirmaier, C.; Laporte, L.; Schenck, C. C.; Holten, D. J. Phys. Chem. 1995, 99, 8903-8909. (c) Kirmaier, C.; Laporte, L.; Schenck, C. C.; Holten, D. J. Phys. Chem. 1995, 99, 89108917. (d) Laporte, L.; Kirmaier, C.; Schenck, C. C.; Holten, D. Chem. Phys. 1995, 197, 8903-8909. (e) Heller, B. A.; Holten, D.; Kirmaier, C. Biochemistry 1996, 35, 15418-15427.

Kirmaier et al. (5) (a) The G(M201)D mutation in Rb. capsulatus and the corresponding G(M203)D mutation in Rb sphaeroides do not affect the redox potential or optical characteristics of P, but only the optical and vibrational properties of BChlL.3a,5b Most importantly, this Asp clearly significantly raises the free energy of state P+BChlL- (probably 100-150 meV) as deduced by effects on electron transfer in a number of Rb. capsulatus RCs containing the G(M201)D mutation.3 The vibrational data indicate that D is not hydrogen bonded to the ring V keto group of BChlL and is most readily interpreted in terms of the D being ionized, at least at room temperature.24 An increase in the free energy of P+BChlL- is consistent with this ionization state of the Asp but also can be accounted for through dipole-dipole or other effects if the Asp is protonated, as long as the effects significantly destabilize P+BChlL-.3a,24 The proposed displacement5c of a nearby water molecule found in the wild-type Rb. sphaeroides1a structure could make such a contribution. Hydrogen bonding involving BChlL with such water molecules will not contribute since such bonding is not commensurate with the vibrational data on wt RCs or the G(M201)D mutant (at least when P is neutral).6,7,22a,24 (b) Williams, J. C.; Alden, R. H.; Murchison, H. A.; Peloquin, J. M.; Woodbury, N. W.; Allen, J. P. Biochemistry 1992, 31, 11029-11037. (c) Fyfe, P. K.; Ridge, J. P.; McAuley, K. E.; Cogdell, R. J.; Isaacs, N. W.; Jones, M. R Biochemistry 2000, 39, 5953-5960. (6) (a) Lutz, M.; Robert, B. In Biological Applications or Raman Spectroscopy; Spiro, T. G., Ed.; Wiley: New York, 1988; Vol 3, pp 347411. (b) Lutz, M.; Mantele, W. In Chlorophylls; Scheer, H., Ed.; CRC Press: Boca Raton, FL, 1991; pp 855-902. (c) Lutz, M. Biospectroscopy 1995, 1, 313-327. (7) Palaniappan, V.; Martin, P. C.; Chynwat, V.; Frank, H. A.; Bocian, D. F. J. Am. Chem. Soc. 1993, 115, 12035-12049. (8) Bylina, E. J.; Kirmaier, C.; McDowell, L.; Holten, D.; Youvan, D. C. Nature 1988, 336, 182-184. (9) (a) Breton, J. Biochim. Biophys. Acta 1985, 819, 235. (b) Breton, J.; Bylina, E. J.; Youvan, D. C. Biochemistry 1989, 28, 6423-6429. (10) Palanniappan, V.; Bocian, D. F. J. Am. Chem. Soc. 1995, 117, 3647-3648. (11) Similar effects to those found on the optical and vibrational properties of BPhL upon the removal of the hydrogen bond to Glu L104 are found upon removal of the analogous hydrogen bond to the BChl (β) that is incorporated into this site in place of BPhL as a result of the “beta” or “H” mutation L(M214)H in Rb. sphaeroides.4b,22a (12) Muh, F.; Williams, J. C.; Allen, J. P.; Lubitz, W. Biochemistry 1998, 37, 13066-13074. (13) (a) Bylina, E. J.; Youvan, D. C. Proc. Natl. Acad. Sci. U.S.A. 1988, 85, 7226-7230. (b) Laible, P. D.; Hanson, D. K. Manuscript in preparation. (c) Kirmaier, C.; Czarnecki, K.; Laible, P. D.; Hata, A. N.; Bocian, D. F.; Holten, D.; Hanson, D. K., in press. See also ref 3d. (14) Heller, B. A.; Holten, D.; Kirmaier, C. Biochemistry 1995, 34, 5294-5302. (15) (a) Kirmaier, C.; Holten, D. Biochemistry 1991, 30, 609-613. (b) Yang, S. I.; Li, J.; Cho, H. S.; Kim, D.; Bocian, D. F.; Holten, D.; Lindsey, J. S. J. Mater. Chem. 2000, 10, 283-296. (16) Palaniappan, V.; Schenck, C. C.; Bocian, D. F. J. Phys. Chem. 1995, 99, 17049-17058. (17) Yu, N.-T.; Srivastava, R. B. J. Raman Spectrosc. 1980, 9, 166171. (18) Diers, J. R.; Bocian, D. F. J. Am. Chem. Soc. 1995, 117, 66296630. (19) Cherepy, J. M.; Shreve, A. P.; Moore, L. J.; Boxer, S. G.; Mathies, R. A. Biochemistry 1997, 36, 8559-8566. (20) Cherepy, N. J.; Shreve, A. P.; Moore, L. J.; Franzen, S.; Boxer, S. G.; Mathies, R. A. J. Phys. Chem. 1994, 98, 6023-6029. (21) Shreve, A. P.; Cherepy, N. J.; Mathies, R. A. Appl. Spectrosc. 1992, 46, 707-711. (22) (a) Czarnecki, K.; Schenck, C. C.; Bocian, D. F. Biochemistry 1997, 36, 14697-14704. (b) Laporte, L. L.; Palaniappan, V.; Davis, D. G.; Kirmaier, C.; Schenck, C. C.; Holten, D.; Bocian, D. F. J. Phys. Chem. 1996, 100, 17696-17707. (c) Czarnecki, K.; Diers, J. R.; Chynwat, V.; Erickson, J. P.; Frank, H. A.; Bocian, D. F. J. Am. Chem. Soc. 1997, 119, 415-426. (d) Czarnecki, K.; Chynwat, V.; Erickson, J. P.; Frank, H. A.; Bocian, D. F. J. Am. Chem. Soc. 1997, 119, 2594-2595. (23) (a) Cherepy, N. J.; Holzwarth, A.; Mathies, R. A. Biochemistry 1995, 34, 5288-5293. (b) Cherepy, N. J.; Shreve, A. P.; Moore, L. J.; Boxer, S. G.; Mathies, R. A. J. Phys. Chem. B 1997, 101, 3250-3260. (24) Czarnecki, K.; Kirmaier, C.; Holten, D.; Bocian, D. F. J. Phys. Chem. A 1999, 103, 2235-2246. (25) Cua, A.; Kirmaier, C.; Holten, D.; Bocian, D. F. Biochemistry 1998, 37, 6394-6401. (26) Because the data extend to only 3.8 ns and this at best is about four 1/e times of the slower component, the error in the associated time constant for the longer-lived component in the anion region and P-bleaching decay is much larger than indicated by simple best fits (1.0 ( 0.4 ns and 1.2 ( 0.4 ns, respectively). For example, in the anion region, holding the slow component fixed at 1.5, 2.0, 2.5, or 3.0 ns gives visually good fits to the gion data in Figure 6, with the value of the faster time constant only

Probing the Bacterial Photosynthetic Reaction Center marginally changed. The use of time constants longer than about 4 ns for the slow component gives poorer fits and unreasonable spectra at the asymptote of the decay. These considerations suggest that the time constant of the slower component is likely in the 1-4 ns range. This behavior parallels that previously observed for the DH and KDH mutants.3a,b (27) As is the case for the DH and KDH mutants, we cannot rule out the presence of a third, small-amplitude component (several percent) to the P-bleaching decay in the V(M131)D-DH RC that might reflect some charge-recombination of the L-side intermediate to the ground-state competing with P+QA- formation (Figure 2B). (28) (a) The amplitude spectra in Figure 7 were generated from fits in which the value of the longer component was held fixed at 2.5 ns. Varying this time constant between 1.0 and 3.5 ns does not appreciably affect the derived spectra. (b) The amplitude spectrum of the slower component is also a second weak feature near 700 nm that is present in P+QA- spectra and that can be assigned to P+.3b,c (29) The vibrations enhanced with QY excitation include the stretching modes of the C10a-carbomethoxy (1725-1755 cm-1) and C9-keto (16851705 cm-1) carbonyl groups (both on ring V), the C2a-acetyl (1650-1680 cm-1) carbonyl group (on ring II), and stretching modes of the CaCm and unsaturated CbCb bonds (1600-1640 cm-1).5d,e,6,7 The remaining stretching modes of the CaCm bonds as well as those of the CaCb and CaN bonds occur at lower frequencies (1300-1600 cm-1; not shown).6,7 (30) The frequencies of the RR bands are of particular interest because they reflect the properties of the ground electronic states of the cofactors. In contrast, the RR intensities are strongly affected by the properties of the QY excited states of the chromophores (via origin shifts and/or dephasing times of certain modes). These particular excited-state properties are outside the scope of this paper. Inspection of the RR data shown in Figures 8 and 9 clearly shows that the RR intensities of the V(M131)D versus wild-type RCs are different at a given excitation wavelength. This observation is similar to that made in previous RR studies of genetically modified RCs.25 These studies have shown that the RR intensities of the cofactors can be significantly affected by altering protein residues in the vicinity of a particular cofactor, even in cases when the genetic modification does not affect the ground-state absorption features or vibrational frequencies of the cofactor to any appreciable extent.25 (31) Peloquin, J. M.; Bylina, E. J.; Youvan, D. C.; Bocian, D. F. Biochemistry 1990, 29, 8417-8424. (32) The same spectral and kinetic data are obtained on the DH mutant in an Rb. capsulatus carotenoidless strain as we have found in the standard carotenoid-containing strain. This finding further rules out any possibility that the 527-nm absorption decrease derives from a carotenoid band shift derived simply from L-side charge separation. We also find that M-side charge separation to form P+BPhM- occurs in the Rb. sphaeroides DH mutant but with a yield of only ∼7%, half that found in the Rb. capsulatus DH mutant.13c (33) (a) Electron transfer to the M-side also has been observed in timeresolved experiments on an Rb. sphaeroides mutant in which BChlM has been replaced by a BPh,34b an Rb. capsulatus mutant involving alterations/ swapping of large sections of the L and M polypeptides,34c and Rb. capsulatus wild-type RCs at low temperature at high excitation intensities.34d (b) Katilius, E.; Turanchik, T.; Lin, S.; Taguchi, A. K. W.; Woodbury, N. W. J. Phys. Chem. B 1999, 103, 7386-7389. (c) Lin, S.; Xiao, W.; Eastman,

J. Phys. Chem. B, Vol. 106, No. 2, 2002 503 J. E.; Taguchi, A. K. W.; Woodbury, N. W. Biochemistry 1996, 35, 31873196. (d) Lin, S.; Jackson, A.; Taguchi, A. K. W.; Woodbury, N. W. J. Phys. Chem. B 1999, 103, 4757-4793. (34) BPhM- has been produced in steady-state photochemical trapping experiments: Robert, B.; Tiede, D. M.; Lutz, M. FEBS Lett. 1985, 183, 326-330. Kellogg, E. C.; Kolaczkowski, S.; Wasiewlewski, M. R.; Tiede, D. M. Photosynth. Res. 1989, 22, 47-59. Gray, K. A.; Wachtveitl, J.; Oesterhelt, D. Eur. J. Biochem. 1992, 207, 723-731. (35) (a) Calculations on hydrogen bonding to the ring V keto group of a BPh36a and experimental results on manipulation of hydrogen bonds to 22b,35b,c P indicate that these hydrogen bonds generally shift the redox properties of the cofactors by 50-80 mV. (b) Allen, J. P.; Williams, J. C. J. Bioenergetics Biomembranes 1995, 27, 275-283. (c) Ivancich, A.; Atrz, K.; Williams, J. C.; Allen, J. P.; Mattioli, T. A. Biochemistry 1998, 37, 11812-11820. (36) For discussions of the factors that may contribute to directionality, see refs 3, 36a-h, and 37. (a) Michel-Beyerle, M. E.; Plato, M.; Deisenhofer, J.; Michel, H.; Bixon, M.; Jortner, J. Biochim. Biophys. Acta 1988, 932, 52-70. (b) Scherer, P. O. J.; Fischer, S. F. Chem. Phys. 1989, 131, 115127 (c) Bixon, M.; Jortner, J.; Michel-Beyerle, M. E. Biochim. Biophys. Acta 1991, 1056, 301-315. (d) Zhang, L. Y.; Friesner, R. A. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 13603-13605. (e) Ivashin, N.; Kallenbring, B.; Larsson, S.; Hansson, O. J. Phys. Chem. B 1998 102, 5017-5022. (f) Hasegawa, J.; Nakatsuji, H. J. Phys. Chem. B 1998 102, 10420-10439. (g) Kolbasov, D.; Scherz, A. J. Phys. Chem. B 2000, 104, 1802-1809. (h) Pudlak, M.; Pincak, R. Chem. Phys. Lett. 2001, 342, 587-592. (37) (a) Parson, W. W.; Chu, Z.-T.; Warshel, A. Biochim. Biophys Acta 1990, 1017, 251-272. (b) Alden, R. G.; Parson, W. W.; Chu, Z. T.; Warshel, A. J. Am. Chem. Soc. 1995, 117, 12284. (c) Gunner, M. R.; Nicholls, A..; Honig, B. J. Phys. Chem. 1996, 100, 4277-4291. (38) (a) Steffen, M. A.; Lao, K.; Boxer S. G. Science 1994, 264, 810816. (b) van Dijk, B.; Hoff, A. J.; Shkuropatov, A. Ya. J. Phys. Chem. 1998, 102, 8091-8099. (39) (a) Thompson, M. A.; Zerner, M. C.; Fajer, J. J. Am. Chem. Soc. 1991, 113, 8210-8215. (b) Marchi, M.; Gehlen, J. N.;, Chandler, D.; Newton, M. Science 1994, 263, 499-502. (c) Blomberg, M. R. A.; Siegbahn, P. E. M.; Babcock, G. T. J. Am. Chem. Soc. 1998, 120, 88128824. (40) (a) Residue L104 is Gln in Chloroflexus aurantiacus RCs1e and in the Rb. capsulatus E(L104)Q mutant.8 In both RCs, the BPhL ground-state QX band (538 nm) and anion band in P+BPhL- (655 nm)40b are blue shifted by a fraction of the magnitudes observed in the Rb. capsulatus Glu(L104) f Leu mutant.8 This comparison suggests that Gln may be hydrogen bonded to the ring V keto group of BPhL in the Chloroflexus aurantiacus RC and the Rb. capsulatus E(L104)Q mutant, but that the hydrogen bond to Gln is not as strong as that to Glu in wild-type Rb. capsulatus (and Rb. sphaeroides).8 (b) Kirmaier, C.; Blankenship, R. E.; Holten, D. Biochim. Biophys. Acta 1986, 850, 275-285. (41) Gouterman, M. In The Porphyrins; Dolphin, D., Ed.; Academic Press: New York, 1978; Vol III, pp 1-165. (42) (a) Petke, J. S.; Maggiora, G. M.; Shipman, L. L.; Christoffersen, R. E. Photochem. Photobiol 1980, 32, 661-667. (b) Petke, J. S.; Maggiora, G. M.; Shipman, L. L.; Christoffersen, R. E. Photochem. Photobiol 1981, 33, 663-671.