Photointermediates of the Rhodopsin S186A Mutant as a Probe of the

Furthermore, the UV−vis absorption maximum of S186A BSI was red-shifted by almost 20 nm relative to that of wild-type BSI, and the deprotonation of ...
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J. Phys. Chem. C 2007, 111, 8843-8848

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Photointermediates of the Rhodopsin S186A Mutant as a Probe of the Hydrogen-Bond Network in the Chromophore Pocket and the Mechanism of Counterion Switch† Elsa C. Y. Yan,*,‡ Jacqueline Epps,§ James W. Lewis,§ Istvan Szundi,§ Aditi Bhagat,‡ Thomas P. Sakmar,‡ and David S. Kliger*,§ Laboratory of Molecular Biology and Biochemistry, Rockefeller UniVersity, 1230 York AVenue, New York, New York 10021, and Department of Chemistry and Biochemistry, UniVersity of California, Santa Cruz, Santa Cruz, California 95064 ReceiVed: October 31, 2006; In Final Form: January 9, 2007

The role of a specific serine located in the retinylidene chromophore-binding pocket of bovine rhodopsin was investigated to determine its role in the mechanism of receptor photoactivaiton. The S186A mutant of rhodopsin was expressed in HEK293S GnTI- cells, and the UV-vis absorbance change of the pigment in n-dodecylβ-D-maltoside detergent was measured as a function of time after photoexcitation with 7 ns laser pulses. Although S186A showed a normal bathorhodopsin (Batho), the microscopic rate constant for the back reaction of S186A blue-shifted intermediate (BSI) to Batho and the forward reaction of S186A BSI to lumirhodopsin (Lumi) were both significantly reduced. Furthermore, the UV-vis absorption maximum of S186A BSI was red-shifted by almost 20 nm relative to that of wild-type BSI, and the deprotonation of the Schiff base was unusually rapid and was complete in microseconds. The observed large mutagenic perturbations to the kinetics and photointermediate spectra suggest that the hydroxyl group of Ser186 interacts with the protonated Schiff base and/or its counterion after photoexcitation. A model is proposed for reorientation of the hydroxyl group of Ser186 upon formation of BSI that is part of a rearrangement of the hydrogen-bond network in the chromophore-binding pocket to facilitate the switch of counterion from Glu113 to Glu181 in the photoactivation process of rhodopsin.

Introduction Photoactivation of Rhodopsin. The rod-cell visual pigment rhodopsin is the only seven-helical G protein-coupled receptor (GPCR) whose crystal structure has been determined.10,15,16,27,28 The transmembrane domain of rhodopsin forms a binding pocket for 11-cis-retinal, which is linked to the opsin protein through a protonated Schiff base (PSB) at Lys296 on helix 7. The positive charge on the PSB is stabilized in the dark by a counterion, Glu113, on transmembrane helix 3. After absorption of a photon, 11-cis-retinal isomerizes to all-trans retinal, and the active state of the receptor is formed after a progression through several intermediates. The primary transient photoproduct, photorhodopsin (Photo), is formed from the ground state within 200 fs after photon absorption.6,22 Relaxation of Photo leads to formation of bathorhodopsin (Batho)5 which decays through the mechanism summarized in Figure 1 to produce the final product metarhodopsin II (Meta II) which activates the G protein transducin (Gt) to trigger the visual signal cascade.1 Counterion Switch. Glu181 on extracellular loop 2 (E2) of rhodopsin was shown to be the primary counterion of the PSB in Meta I, implying that the counterion switches from Glu113 in the dark state to Glu181 in the Meta I state after photoexcitation.30 The E181Q mutant was prepared in digitonin †

Part of the special issue “Kenneth B. Eisenthal Festschrift”. * Corresponding authors. Phone: (212) 327-8284 (E.C.Y.Y.); (831) 4592106 (D.S.K.). Fax: (212) 327-7904 (E.C.Y.Y.); (831) 459-2935 (D.S.K.). E-mail: [email protected] (E.C.Y.Y.); [email protected] (D.S.K.). ‡ Rockefeller University. § University of California, Santa Cruz.

Figure 1. Rhodopsin (Rho) bleaching sequence near physiological temperatures in the detergent DM. Some of the intermediates appearing above can be trapped after low-temperature photolysis, but those shown in italics, BSI (whose equilibrated mixture with Batho is sometimes called BL) and Meta I380, only build up appreciable concentrations near physiological temperatures (refs 4, 8, and 25) or in the case of Photo, cannot be trapped because it results from transient localized heating. The time constants given are appropriate for DM suspensions of rhodopsin near 20 °C. This general scheme also holds for membrane samples with the same time constants up to Lumi II formation. However, at lower temperatures in membrane and in the detergent digitonin the Lumi II a Meta I380 equilibrium is back-shifted and the branch shown in curly brackets above becomes appreciable leading to the appearance of a Meta I480 a Meta II equilibrium which is not seen in the detergent DM. Low temperatures also back-shift the Meta I480 a Meta II equilibrium allowing Meta I480 to be trapped. The trapped form is often referred to as metarhodopsin I (Meta I). The above scheme is somewhat simplified in that a single Meta II product is shown. This neglects proton uptake that occurs between two isochromic forms of Meta II to produce the G protein activating form (sometimes called R*). In DM suspensions this proton uptake is completely forward shifted.

suspension, and its Meta I state was trapped using low temperature (4 °C). The pKa of the PSB in the Meta I state was

10.1021/jp067172o CCC: $37.00 © 2007 American Chemical Society Published on Web 02/22/2007

8844 J. Phys. Chem. C, Vol. 111, No. 25, 2007 found to change from >9 for wild-type rhodopsin to ∼6 for E181Q. This behavior is very similar to the titration of E113Q in the dark state, which was used to conclude that Glu113 is the counterion for the PSB in the dark.21,32 It was argued that the E181Q mutation removes the counterion in the Meta I state and changes the electrostatic environment of the PSB, leading to a shift of pKa. The role of Glu181 was also investigated by Fourier transform infrared (FTIR)11 and transient UV-vis absorption spectroscopy.9 The vibrational difference spectra of E181Q in the Meta I and Meta II states showed that the Meta I conformation in the E181Q mutant adopts a conformation that is Meta II-like, suggesting that the removal of the native charge on Glu181 significantly changes the structure of Meta I and perturbs the Meta I/Meta II equilibrium.11 The results also showed that the Schiff base is partly deprotonated in Meta I, supporting the idea that Glu181 is the primary counterion in the Meta I state. Using transient UV-vis absorption spectroscopy, we studied the effect of the Glu181 mutations on the kinetics of rhodopsin activation.9 We found that the kinetics and spectra of intermediates in the photoactivation pathway were highly perturbed by the Glu181 mutations studied. In the case of E181F, early deprotonation of Lumi II, the photointermediate which immediately precedes the Meta I intermediates, was induced. Thus, the removal of the counterion of the PSB in Meta I by the Glu181 mutations has a large energetic and structural impact on the overall photoactivation process. This reinforces the role of Glu181 as the counterion of the PSB in Meta I. Hydrogen Bonds and Photoactivation. The counterion switch process is likely coupled to the structural evolution of a hydrogen-bond network in rhodopsin. Previous vibrational studies have shown that the strength of the PSB hydrogen bond changes continuously along the photoactivation pathway.17,23 These changes should be correlated with the counterion switch process because strong electrostatic interaction of the PSB is shifted from Glu113 to Glu181. Thus, further investigation of how the hydrogen-bond network couples to the counterion switch process, in the temporal middle of the photoactivation pathway (late microsecond range),3,4,8,26 should be important to understanding the rhodopsin signaling process. Ser186 is a key residue of the hydrogen-bond network in the retinal binding pocket (Figure 2). It is situated between Glu113 and Glu181. The backbone oxygen is linked to the backbone nitrogen of Glu181, and the hydroxyl group of Ser186 is hydrogen bonded to two water molecules in the binding pocket. Ser186 is close to the PSB-counterion system but appears to be too far away (∼4 Å) to directly form a hydrogen bond with either the PSB or counterion before isomerization. Since hydrogen bonds have an unusually large role in stabilizing structures in the nonpolar transmembrane region in the dark state, it seems likely that specific hydrogen-bonded patterns may be characteristic features of rhodopsin’s photointermediates. In the dark, the side chain of Ser186 is close to, but not yet hydrogen bonded, with the PSB-counterion complex, the most polar part of rhodopsin’s transmembrane region. Is it possible that the hydroxyl group of Ser186 will reorganize in the photoactivation process? Which photointermediates does the polar character of Ser186 affect? To answer these questions, we have investigated the effect of removing the Ser186 hydroxyl group on photointermediate spectra and kinetics during the activation process of rhodopsin using transient UV-vis absorption spectroscopy. Materials and Methods Stably Transfected Cell Line. Expression of the S186A mutant opsin was carried out in suspension culture of the

Yan et al.

Figure 2. Hydrogen-bond network in the binding pocket of rhodopsin. Solid lines show distances ranging from 2.6 (Glu181-W6) to 3.2 Å (PSB-Glu113) representing possible hydrogen bonds in rhodopsin. The dashed line shows the distance between the side chain of Ser186 and Glu113 (4.3 Å) which exceeds hydrogen-bond lengths but could be significantly reduced by rotation of the Ser186 side chain in one of the photointermediates. This model and numbering scheme for the water molecules (W6, W16, and W18) are adapted from Li et al. (ref 10).

HEK293S GnTI- system developed by Reeves and co-workers.19,20 The HEK293S GnTI- cells and pACMV-tetO expression vector were obtained as a gift from Dr. Reeves (Essex, U.K.). To avoid recurring costly transient transfection and to increase the expression yield, the cell line stably transfected with the S186A mutant bovine opsin gene was made based on the method slightly modified from the one described previously.19,20 To prepare the DNA construct for the transfection, site-directed mutagenesis of bovine opsin was first carried out in the expression vector pMT4 as previously reported.29 The S186A mutant gene was PCR amplified by a pair of primers that engineer a KpnI site at the 5′ end and a NotI site at the 3′ end. The S186A mutant opsin gene was then subcloned to the pACMV-tetO vector with the KpnI/NotI restriction sites. The sequence of the construct was confirmed by DNA sequencing. To make the stable cell line, on day 1 the HEK293S (GnTI-) cells were transfected with the DNA construct using Lipofatamine (Invitrogen). The transfected cells were split and diluted into different ratios (1:5-100) on day 2. They were maintained in medium 1:1 DMEM/F12 supplemented with FBS (10%), penicillin G (100 units/mL), streptomycin (100 µg/mL), and glutamine (300 µg/mL). The antibiotic Geneticin (300 µg/mL) was present in the medium for selection. In about 2 weeks, Geneticin-resistant colonies that were stably transfected with the expression vector were formed. Ten clones were isolated using cloning rings. The clones were propagated via expansion onto a 10 cm tissue culture plate to screen for the highest expression. To assess the expression level of each clone, opsin expression was induced by adding tetracycline (2 µg/mL) and sodium butyrate (5 mM). After 48 h, the cells were harvested. They were then regenerated with 11-cis-retinal for 3 h and solubilized with 1% (w/v) n-dodecyl-β-D-maltoside (DM) for another 3 h. Half of the cell lysate was collected for modified Lowry assay to measure the amount of total solubilized protein for use as a normalization factor to compare expression level. The rest of the cell lysate was spun at 60 000 rpm for 30 min.

Photointermediates of the Rhodopsin S186A Mutant UV-vis spectra of the supernatant were taken before and after the samples were bleached with light λ > 495 nm. The intensity of the 500 nm peak in the difference spectrum for each clone was normalized to the amount of total solubilized protein. The clone that gave the largest normalized change at 500 nm was further propagated, then frozen and stored in liquid nitrogen until required for protein expression. Preparation of Rhodopsin Mutant. Protein expression was carried out in 250 mL of suspension culture grown in a 1 L spinner flask at 37 °C and 5% CO2. The growth medium recipe was adapted from Reeves and co-workers19,20 containing calcium-free DMEM medium supplemented with FBS (10%), Primatone RL-UF (0.3% w/v), pluronic acid (0.1% v/v), and dextran sulfate (300 µg/mL). For inoculation, the stable cell line was first grown to confluent on 10 cm tissue culture dishes. On day 1, the cells were trypsinized and inoculated to 200 mL of medium in the spinner flask at a density of 0.5 × 106 cells/ mL. Fifty milliliters of fresh growth medium was added to the spinner flask on day 3. On day 5, the culture was supplemented with 2.5 mL of glucose (20% w/v) and 7.5 mL of Primatone RL-UF (10% w/v). Tetracycline (2 µg/mL) and sodium butyrate (5 mM) were added on day 6 for inducing protein expression. The cells were then harvested on day 8 at a cell density of 3 × 106 cells/mL. The cells were regenerated with 11-cis-retinal, and the pigment was purified by an immuno-affinity method as described previously.12,13 The final elution buffer contained 0.1% DM, 10 mM Tris, 30 mM NaCl, 60 mM KCl, and 2 mM MgCl2. A UV-vis spectrum of the elution was taken to measure the concentration of the S186A pigment (Perkin-Elmer Lambda800). The sample was then concentrated by a filter device (Amicon) to ∼25 µM (OD500 ∼ 1) for the time-resolved measurements. Each batch of pigment was diluted to 450 µL (the capacity of the syringe pump on the time-resolved absorbance apparatus) with the final elution buffer and degassed for 90 s at room temperature under house vacuum to prevent bubble formation during data collection. Samples were maintained at 20 °C during measurements. Collection of Time-Resolved Absorbance Difference Spectra. Suspensions were pumped in 1 µL aliquots into the cuvette before being photoexcited by 7 ns (full width at half-maximum) pulses from a dye laser that produced vertically polarized light at 477 nm. The energy delivered to the sample was 80-100 µJ/mm2. Alterations in the absorption spectrum monitored at time delays ranging from 30 ns to 1 ms after photolysis were measured using a gated optical array detector as previously described8 except that an Andor DH-520 intensified CCD detector replaced the PAR 1420 intensified photodiode array. The ability of the Andor system to collect I(λ, 0) (the probe intensity before laser photolysis) and I(λ, t) (the probe intensity at the time, t, after photolysis) on the same aliquot of sample reduced consumption of mutant pigments by a factor of 2 over what had been previously possible using the PAR 1420 system, improving the overall signal-to-noise ratio. The probe source polarization axis was set at 54.7° relative to the laser polarization direction to avoid kinetic artifacts due to rotational diffusion. The pathlengths of the actinic light and probe light in the sample were 0.5 and 2 mm, respectively, and the laser and probe beams crossed at a right angle. Data Analysis. The experimental difference spectra, {∆A(λ, t)}, were analyzed using singular value decomposition (svd) followed by global exponential fitting.2 The purpose of svd is to reduce the noise level in the data so that the nonlinear leastsquares process used to find the lifetimes (global exponential fitting) proceeds more efficiently. In svd the data matrix, ∆A,

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Figure 3. Time-resolved absorbance changes observed at early times after 20 °C photolysis of the S186A mutant of rhodopsin. Solid lines in descending thicknesses show absorbance difference spectra collected at 30, 60, 120, and 240 ns after photolysis, and dotted lines in descending sizes continue the time delays to 480 ns, 1, 2, and 4 µs. The increase in absorbance at 30 ns, peaking near 560 nm, is due to formation of the S186A Batho intermediate which decays on the time scale shown to produce blue-shifted products.

is split into a product of three matrices: ∆A ) U*S*V′, where U is a matrix of orthogonal spectral vectors, V is their time dependence, and S contains the significance values indicative of the contributions of the U and V vectors to the experimental data. The significant vectors in the temporal matrix, V, were fitted to a sum of exponential functions followed by calculation of the spectral amplitudes, the b-spectra. From the b-spectra and exponential functions the matrix of reproduced data, ∆a, was calculated:

∆a(λ, t) ≡ b0(λ) + b1(λ) exp(-t/τ1) + b2(λ) exp(-t/τ2) + ... where the τi are the apparent lifetimes and the bi(λ) are b-spectra corresponding to difference spectra which decay with the associated lifetime. The fit produced a best estimate of what the true difference spectra were at each delay time. The difference between the experimental data matrix and the reproduced data, R ) ∆A - ∆a, gives the matrix of residuals that was used to judge the quality of the fit. The residual matrix of an adequate fit should contain only featureless noise caused by the finite number of photons detected in each measurement. Spectra of intermediates were determined from b-spectra as described previously.24 Results Time-resolved absorbance difference spectra collected at delays from 30 ns to 4 µs after photoexcitation of the S186A mutant of rhodopsin are shown in Figure 3. The positive absorbance change to the red of about 520 nm detected at 30 ns is clear evidence for the formation of the S186A Batho photointermediate. The decay of that absorbance on the tens of nanoseconds time scale demonstrates that S186A Batho has stability similar to that of wild-type Batho. Although the data in Figure 3 from 480 ns to 4 µs (the dotted curves) are noisy, a continuing trend can be seen. Analysis shows that significant further decay of S186A Batho takes place on that time scale to produce S186A Lumi. The S186A Lumi formation process is noticeably slower than wild-type Lumi formation, which is 95% complete within 500 ns at 20 °C.2 Further absorbance changes detected up to 1 ms, the time scale for decay of S186A Lumi, are shown on an expanded scale in Figure 4, beginning with the last two curves from Figure 3. That data show a visible absorbance (characteristic of PSB intermediates) decaying as

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Figure 4. Time-resolved absorbance changes detected at late times after 20 °C photolysis of the S186A mutant of rhodopsin. Solid lines show absorbance difference spectra collected at 2 (heavy line), 4, 10, 20, 50, 200 µs, and 1ms (heavy line) after photolysis. During this time period the protonated Schiff base deprotonates.

absorbance in the 380 nm region (characteristic of deprotonated SB intermediates) grows. The data in Figures 3 and 4 were best fit by three exponential processes with time constants 55 ns, 1.1 µs, and 35 µs corresponding to the b-spectra shown by the data points in the upper panel of Figure 5. The residuals characterizing that fit are shown in the lower panel of Figure 5. The first two timedependent b-spectra have positive absorbance changes in the red part of the spectrum which denotes that S186A Batho decays in both these processes. The amplitude of the second of these (lifetime ) 1.1 µs) is much smaller than the first one (lifetime ) 55 ns). This behavior differs significantly from what is seen after photoexcitation of wild-type rhodopsin, where at 20 °C Batho decays in two approximately equal amplitude processes because the Batho a BSI equilibrium constant is near 1.2 A further difference from wild-type rhodopsin is that the final formation of the S186A deprotonated SB intermediates is at least an order of magnitude faster.3 The b-spectra shown in Figure 5 could be fit to a mechanism similar to that shown in Figure 1 but with some of the microscopic rate constants and photointermediate λmax values significantly perturbed. The agreement of the fit (lines) with the experimental b-spectra (points) is shown in the upper panel of Figure 5, and the spectra of the model photointermediates used in the fit are shown in Figure 6 (also plotted against the experimental data). The λmax’s of the S186A photointermediates are given in Table 1, and the microscopic rate constants determined from the fit are given in Table 2. While it is clear that the SB deprotonates fully in the final S186A product, the fact that the 380 nm absorbance forms in a process described by a single exponential makes it difficult to identify unambiguously which of the several identified deprotonated SB intermediates is responsible or whether the final product is a mixture of more than one (see below). Discussion It has long been known that the functionally important absorbance of visual pigments arises from a protonated retinylidene Schiff base which is stabilized by a nearby carboxylate counterion. Such a polar environment seems incompatible with the otherwise hydrophobic character of the transmembrane region characteristic of 7-transmembrane receptors where it occurs, but the relatively nonpolar chromophore pocket could explain why specific polar amino acid side chains contribute to tuning the λmax of visual pigment absorbances,14,33,34 accounting

Figure 5. b-Spectra and residuals from a three-exponential fit of timeresolved absorbance changes in the S186A mutant of rhodopsin. Points in the top panel show the shapes of the three time-dependent b-spectra (labeled with the observed lifetimes) and the time-independent bspectrum, b0, obtained by fitting the S186A data. Points in the lower panel show the residuals obtained after subtracting the best threeexponential fit from the data. The points at the bottom of the lower panel correspond to residuals at 30 ns, and residuals at successive times are offset up by 0.002 absorbance units. Fits using fewer exponentials showed significant curvature in the residuals at some of the delay times. No significant improvement in the flatness of the above residuals was obtained by using more exponential terms. Smooth curves in the upper panel show the b-spectra modeled using the modified rhodopsin mechanism and photointermediates (see text). In that mechanism the small amplitude of the second time-dependent b-spectrum (0) relative to that of the first (4) results from an unusual forward shift of the S186A Batho-S186A BSI equilibrium.

Figure 6. Spectra of S186A photointermediates. Smooth curves show the model spectra of photointermediates used in fitting the b-spectra. Points show the results at each wavelength for spectral changes associated with each photointermediate (S186 Batho, 4; S186A BSI, 0; S186A Lumi, 3; 380 nm absorbing product, O).

for the wide variety of visual pigments found in nature. Also surprising for a hydrophobic region, rhodopsin crystal structures and subsequent molecular dynamic simulations showed water molecules in the transmembrane portion.10,15,16,27,35 Current models suggest these water molecules interact with polar amino acid side chains in a network of hydrogen bonds that undergoes

Photointermediates of the Rhodopsin S186A Mutant

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TABLE 1: Absorbance Maxima of Photointermediates S186A mutant rhodopsin a

Batho

BSI

Lumi

product

526 nm 529 nma

496 nm 477 nma

494 nm 492 nma

384 nm 380 nm

Ref 2.

TABLE 2: Microscopic Rate Constants Batho f BSI

Batho r BSI

BSI f Lumi

Lumi f 380 product

S186A mutant 1.7 × 107 s-1 1.5 × 106 s-1 1.0 × 106 s-1 2.9 × 104 s-1 rhodopsin 1.3 × 107 s-1 9.0 × 106 s-1 7.0 × 106 s-1 0.2 × 104 s-1

rearrangement during activation, specifically mediating processes such as the counterion shift which occurs at Meta I.30 The current work investigates the consequences for the rhodopsin photointermediates when the hydroxyl group of Ser186, which is close to the PSB counterion region and hydrogen bonded to two water molecules, is removed by the S186A mutation. The observations of only mildly altered absorption maximum and decay kinetics of S186A Batho suggest the packing of the retinylidene binding pocket has not been significantly perturbed in the Batho state. Previous studies have shown that the large red-shift of visible absorption38 and the unique decoupled hydrogen out-of-plane vibration modes at the C11dC12 bond37 arise from a highly specific distortion along the ethylenic chain of the chromophore.36,39 This distortion is controlled by packing forces that are apparently unaffected by the S186A mutation. This may also imply that the two water molecules (W6 and W18 as shown in Figure 2, numbering scheme adapted from Li et al.10) in the binding pocket are also present in the mutant. One of the water molecules (W6 in Figure 2) is in the path of the 13-methyl of the retinylidene chromophore in the photoisomerization process and is believed to be an important factor contributing to Batho stability.7 If W6 were absent or perturbed by the absence of its partner (W18), it is likely that S186A Batho would decay on the subnanosecond time scale as it does after photoexcitation of the artificial pigment 13-demethylrhodopsin.18 Nonetheless, the removal of the hydroxyl group at the 186 position should affect the positions and orientations of the two water molecules and thereby mechanistically affect the kinetics of later photointermediate decay as discussed below. The perturbation of the S186A mutation to BSI is much more significant than that to Batho. The absorption maximum of S186A BSI is red-shifted by 19 nm relative to that of the wild type. The microscopic rate constants for the back reaction of S186A BSI to S186A Batho and the forward reaction to S186A Lumi are both considerably reduced compared with those of wild-type BSI. This suggests that for the first time at BSI, the hydrogen-bonding character of Ser186 may become important. We propose a model in which a specific rotamer of the side chain of S186A in BSI may be required for both the forward reaction to Lumi and the backward reaction to Batho. Within this model, the removal of a hydroxyl group from Ser186 could introduce unproductive conformations of the two bound waters and/or the PSB and counterion that would lower both the forward and backward rates as observed. It may also account for the significant red-shift of S186A BSI absorbance because a more polar environment blue-shifts the absorbance of PSBs. Support for a change in the environment of the PSB at BSI comes from time-resolved resonance Raman measurements, which have shown that the PSB-counterion hydrogen bond weakens at BSI.17 Given the large perturbation of the BSI λmax and the lack of perturbation of the S186A Lumi spectrum, apparently whatever

specific hydrogen-bonding interaction occurs at BSI between S186 and the PSB is transient. As time proceeds, the chromophore and pocket evolve significantly beyond the dark structure, and with W6 and W18 (Figure 2) nearby, there are numerous hydrogen-bonding possibilities besides Ser186 for the PSB and counterion. Study of more rhodopsin mutants should help to resolve the specific groups involved, and this task will be aided by better structural studies of the late photointermediates. A final unusual property of the S186A mutant is the rapid deprotonation of the PSB. Because several isospectral intermediates absorbing near 380 nm occur after photolysis of wildtype rhodopsin, it is difficult to determine which is formed in S186A. In pigments where 380 nm absorbance develops in two exponential processes, it can be interpreted as Lumi II initially approaching equilibrium with Meta I380 followed by irreversible formation of Meta II on a longer time scale. This does not seem to be the case here, which could indicate that the S186A mutation completely forward shifts the Lumi II a Meta I380 equilibrium. However, 380 nm absorbance forms very quickly after S186A photoexcitation and could occur as early as the time scale associated with Lumi II formation itself. This may indicate that the S186A mutation has a dramatic effect on the pKa of the PSB of Lumi II, as has been seen for the E181F rhodopsin mutant. Further study of mutants connected to these important residues for the counterion shift mechanism should help to determine whether this is the case. Our results reveal an important role of Ser186 in the photoactivation process of rhodopsin. The photokinetics of rhodopsin activation is significantly perturbed by the removal of the hydroxyl group on Ser186. The effect starts as early as the formation of BSI on the tens of nanoseconds time scale. The perturbation persists throughout the activation process as shown by the observed early deprotonation of the Schiff base. The observed spectral shift of S186A BSI (19 nm relative to wild-type BSI) suggests that at BSI the hydroxyl group of Ser186 reorients to interact with the PSB-counterion complex because such a large spectral shift in the mutant is most likely to be associated with the removal of interaction between the PSB-counterion complex and hydroxyl group on Ser186. This reorientation of the hydroxyl group on Ser186 could be a crucial event in the rearrangement of the hydrogen-bond network in the retinal binding pocket upon photoactivation. The Ser186 reorientation proposed by our model is likely to occur in parallel with the changes of hydrogen-bond environment of the PSB in the photoactivation process because the time scales of both processes are similar. Previous vibrational studies by both FTIR and Raman spectroscopies show a dramatic change of the hydrogen-bond environment at the PSB after photoactivation.17,23 Pan et al.17 performed transient Raman studies and measured the CdN stretching frequency (ν) of the PSB in H2O and D2O. They observed a gradual decrease of isotopic shift from Rho to BSI, followed by a larger decrease from BSI to Lumi and then an increase again from Lumi to Meta I. Because the isotopic shift reflects the strength of the hydrogen bond of the proton at the PSB,17,23 the observed change of isotopic shift implies a severe modification of the hydrogenbonding environment at the PSB along the photoactivation pathway, and the modification may involve establishing a new interaction between the hydroxyl group of Ser186 and the PSBcounterion complex. The reorientation of the hydroxyl group of Ser186 is likely to couple to the counterion switch process. Ser186 is situated between Glu113 and Glu181 and is hydrogen bonded to Glu181

8848 J. Phys. Chem. C, Vol. 111, No. 25, 2007 and two water molecules in the binding pocket (Figure 2). Switching the primary electrostatic interaction of the PSB from Glu113 to Glu181 should involve some temporary interaction of the PSB and Ser186. Our results suggest that the PSB and the hydroxyl group of Ser186 start interacting with each other upon formation of BSI. We propose a model wherein the reorientation of the hydroxyl group of Ser186 is part of the rearrangement of the hydrogen-bond network in the retinal binding pocket that assists the switch of counterion from Glu113 to Glu181. Because an electrostatic interaction is the strongest noncovalent force in a hydrophobic environment within proteins, the structural changes driven by the counterion switch process could be a key molecular event that transmits the visual signal from the highly twisted Batho chromophore to the cytoplasmic surface to achieve the active conformation. This molecular picture agrees with the observation that switching ionic interactions in 7-transmembrane GPCRs could be an important molecular mechanism in the activation process.31 Acknowledgment. This research was supported by Grants (EY00983) from the National Institutes of Health (to D.S.K.) and from the Allene Reus Memorial Trust and the Ellisan Medical Foundation (to T.P.S.). We thank Dr. P. Sachdev for advice and assistance in establishing the stable cell lines. References and Notes (1) Hofmann, K. P.; Ja¨ger, S.; Ernst, O. P. Isr. J. Chem. 1995, 35, 339-355. (2) Hug, S. J.; Lewis, J. W.; Einterz, C. M.; Thorgeirsson, T. E.; Kliger, D. S. Biochemistry 1990, 29, 1475-1485. (3) Ja¨ger, S.; Lewis, J. W.; Zvyaga, T. A.; Szundi, I.; Sakmar, T. P.; Kliger, D. S. Biochemistry 1997, 36, 1999-2009. (4) Ja¨ger, S.; Szundi, I.; Lewis, J. W.; Mah, T. L.; Kliger, D. S. Biochemistry 1998, 37, 6998-7005. (5) Kim, J. E.; Mathies, R. A. J. Phys. Chem. A 2002, 106, 85088515. (6) Kukura, P.; McCamant, D. W.; Yoon, S.; Wandschneider, D. B.; Mathies, R. A. Science 2005, 310, 1006-1009. (7) Lewis, J. W.; Fan, G. B.; Sheves, M.; Szundi, I.; Kliger, D. S. J. Am. Chem. Soc. 2001, 123, 10024-10029. (8) Lewis, J. W.; Kliger, D. S. Methods Enzymol. 2000, 315, 164. (9) Lewis, J. W.; Szundi, I.; Kazmi, M. A.; Sakmar, T. P.; Kliger, D. S. Biochemistry 2004, 43, 12614-12621. (10) Li, J.; Edwards, P. C.; Burghammer, M.; Villa, C.; Schertler, G. F. X. J. Mol. Biol. 2004, 343, 1409-1438. (11) Ludeke, S.; Beck, R.; Yan, E. C. Y.; Sakmar, T. P.; Siebert, F.; Vogel, R. J. Mol. Biol. 2005, 353, 345-356. (12) Marin, E. P.; Krishna, A. G.; Archambault, V.; Simuni, E.; Fu, W. Y.; Sakmar, T. P. J. Biol. Chem. 2001, 276, 23873-23880.

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