Photocycle Dynamics in Bacteriorhodopsin in an Ethanol Perturbed

We examine the dependence of the native bacteriorhodopsin (bR) photocycle on ethanol, a solvent thought to perturb lipid structure, and monitor protei...
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12906

J. Phys. Chem. B 2001, 105, 12906-12912

Photocycle Dynamics in Bacteriorhodopsin in an Ethanol Perturbed Lipid Environment Using Time-Resolved Infrared Spectroscopy Tina M. Masciangioli† and Jane K. Rice* NaVal Research Laboratory, Code 6111, Washington, DC, 20375-5342 ReceiVed: June 13, 2001; In Final Form: October 5, 2001

We examine the dependence of the native bacteriorhodopsin (bR) photocycle on ethanol, a solvent thought to perturb lipid structure, and monitor protein and chromophore absorption changes using time-resolved Fourier transform infrared (FTIR) spectroscopy. Ethanol treated samples under two pH conditions, 7 and 8, and two temperatures are examined. A faster M to N transition and slower N to O/bR transition are observed with 3 M ethanol at pH 8. These rates are determined by monitoring several different infrared bands associated with the bR proton pump. An examination of the positions of the infrared bands associated with Asp85 and Asp96 (crucial bR proton transport amino acids) indicates that ethanol produces no obvious differences attributable to bR protein structural changes. Lipid-specific infrared bands in the CH stretching region and methylene region in the bR-lipid system and a model diphytanyl lipid undergo shifts in the presence of ethanol consistent with conformational changes in the lipid acyl chain. It also appears that bR protein infrared bands not associated with the major proton pumping sites are shifted by the presence of ethanol; however, further studies are needed.

Introduction Lipids are intimately involved in the hydrogen-bonding environment of integral membrane proteins. Membrane proteins play an important role in many biological systems, such as photosynthetic reaction centers, ion-gated channels, and the proton pump bacteriorhodopsin (bR), but the mechanism by which the membrane influences the protein function is still unclear. Because bR is well characterized, naturally exists in a lipid environment, and has robust physical properties in comparison with other biological samples, it is an ideal model system for studying lipid-protein interactions and the mechanism of proton transport through membrane proteins.1 In addition, bR is remarkably similar to the visual receptor rhodopsin, which is a member of the G protein-coupled receptors.2 Ligand-gated channels such as rhodopsin,3 which appear to be highly coupled to the lipid membrane, have been used to elucidate the mechanistic action of alcohols and general anesthetics. Bacteriorhodopsin is a protein that exists as part of what is called the purple membrane (PM) of Halobacterium salinarium,4-6 an Archaeo-bacteria (Archaea). Lipids are known to play a significant role in the photocycle dynamics of the proton pump in bacteriorhodopsin. Alteration to the lipid environment by changes to its composition or bonding structure can significantly increase or decrease the rate at which various photocycle intermediates form and decay. The membrane lipids of the Archaea are unique and distinct from those encountered in Eukarya and Bacteria. The PM consists of 25% lipids, of which 60% are phospholipids, 30% are glycolipid, and 10% are neutral lipids, primarily squalenes (SQ).7,8 The methyl ether derivative of phosphatidylglycerophosphate (PGP) is the major phospho* To whom correspondence should be addressed. E-mail: rice@ ccs.nrl.navy.mil. † National Research Council-Naval Research Laboratory Research Associate. E-mail: [email protected].

lipid present in PM. The polar lipids are further distinguished by the fact that the sn-2,3 carbons of the glycerol backbone are attached by ether linkages to two phytanyl chains (saturated palmityl chains with branched methyl groups every four carbons) with typical chain lengths of 20 or 40 carbons.9 The headgroup spacings in phytanyl chain lipids (with branched methyl groups) are larger than the spacings in lipids with saturated, unbranched, alkyl chains.10 It has also been shown that a crucial interaction occurs between bR and a squalene-PGP complex that may be essential for normal bR photocycle activity.11 The proton pumping mechanism of bR can be simplified to three steps. The first is the proton transfer from retinal to the Asp85 amino acid residue during M-intermediate formation,12-14 which results in a proton release to the extracellular (EC) side of the membrane. The formation of M occurs in ∼60 µs. For the cycle to be restored, the second step is a reprotonation of the Schiff base from Asp96 (located on the cytoplasmic (CP) side of retinal) as the N-intermediate forms.15,16 The formation of the N intermediate occurs in ∼1 ms. The Asp85 residue remains protonated in N, Asp96 becomes deprotonated, and the retinal-binding site is then relaxed.17-21 Thermal reisomerization to all-trans retinal is made possible during N because the Schiff base has been reprotonated, lowering the barrier to double-bond rotation.22 The third step is the restoration of a proton to Asp96 from the CP surface of the membrane (∼6 Å distance), during the transition from the N back to the ground-state bR. This transition occurs in ∼6 ms. These changes can be followed in detail by using time-resolved FTIR spectroscopy. Fukuda and Kouyama23 investigated the photocycle of bR using transient visible spectroscopy in the presence of several small alcohols. The technique they used monitored changes in the visible absorption of the covalently attached bR chromophore upon photoinitiation. They found that alcohols increased the rate of N formation and decreased the rate of N decay of the bR photocycle.

10.1021/jp012264t CCC: $20.00 © 2001 American Chemical Society Published on Web 12/05/2001

Dynamics in Bacteriorhodopsin with Ethanol Other evidence of the influence of alcohols on lipids and bR has been reported. Chiou,24 and references therein have shown that lipid structures are dehydrated by alcohols, using FTIR spectroscopy on a model lipid system in reverse micelles. Alcohols have also been shown to affect the hydrogen bonding between helix coils in bR.25 Measurements of the molecular motion of retinal in bR have revealed a wavelength-dependent anisotropy, whereas measurements in bR without ethanol do not. Ethanol was also seen to facilitate the rotation of nonexcited protein, but it did not significantly affect the motion of bR photointermediates. 26 Here, we examine the dependence of the photocycle dynamics during proton pumping of bR with and without ethanol using time-resolved Fourier transform infrared spectroscopy (FTIR). FTIR spectroscopy provides information at the molecular level and allows us to probe structural changes of both the retinal chromophore and protein moiety of bR. We address how ethanol interacts with bR and its lipid membrane, the bR proton pumping kinetics, and the possible mechanisms by which ethanol effects the photocycle of bR. Experimental Section Sample Preparation. Purified purple membrane, isolated from Halobacterium salinarium (S9), was obtained from Professor Mostafa A El-Sayed and Dr. Jianping Wang in the Chemistry Department of Georgia Institute of Technology in Atlanta, GA. A 0.5 mL aliquot of a 0.5 mg/mL solution of bR in deionized water was allowed to dry on a 25 mm × 2 mm CaF2 plate. The dried film was then rehydrated with a phosphate buffer (0.1 M NaCl, 10 mM KH2PO4, various pH values) or the same buffer solution containing 3 M absolute ethanol. The film was immersed in the buffer solution for 1 h and then the excess solution was removed. The hydrated film was then squeezed between two CaF2 plates separated by a 12 µm Teflon spacer. Data were collected from multiple spots on the sample by rotating the sample holder. This sample preparation method was also used for static FTIR measurements which were used to monitor lipid structure in bR. The model lipid diphytanyl phosphatidyl choline (DOPhPC: 1,2-Di-O-phytanyl-sn-glycero-3-phosphocholine), stored in chloroform, was obtained from Avanti Polar Lipids (Alabaster, AL). To obtain the lipid spectra in Figures 3 and 4, the DOPhPC/ chloroform solution was left to dry on a CaF2 plate. Buffer solution was added to the dry film and then the hydrated sample was covered by a second CaF2 plate using a 6 µm spacer. The static second derivative spectra of bR and the model lipid in Figures 3 and 4 were collected at 2 cm-1 resolution using the DTGS detector. Laser Excitation and Time-Resolved FTIR Measurements. The photocycle of bR was initiated using 10 ns, 3-5 mJ/pulse of the second harmonic frequency (532 nm) of a nanosecond Nd:YAG laser (Quantel Model YG581C). The laser was externally triggered at 10 Hz in order to obtain baseline singlebeam data points prior to excitation. To do this, we programmed a digital pulse/delay generator (DG535, Stanford Research) and home-built pulse amplifier that triggered the laser as well as collection of the transient FTIR signal described below. A Bruker IFS/66 FTIR step-scan spectrometer was used for time-resolved FTIR measurements of the modified bR samples. It consisted of a Pentium 100 MHz computer with Bruker OPUS software (OS/2 v.3.0.4), and the internal 16-bit analog to digital converter (ADC) capable of 5 µs resolution. An MCT detector (KMPV11-0.5 J2, Kolmar Technologies, Newburyport, MA) was used. An infrared filter passing 2500 to 850 cm-1 (OCLI

J. Phys. Chem. B, Vol. 105, No. 51, 2001 12907

Figure 1. Temporal evolution of the FTIR difference absorption spectrum in the 1800 to 1720 cm-1 region after laser excitation of bR, in the absence (top) and presence (bottom) of 3 M ethanol (25 °C, pH 8). Spectral traces are shown at 100 µs, 1, 5, and 9 ms. The major absorption band observed is identified as the CdO stretching mode of Asp85 and appears during M-intermediate formation. It shifts to 1755 cm-1 during formation of N. Vertical lines indicate peak maxima in the bR spectrum.

Figure 2. Fast rise and subsequent decay of the N intermediate to bR from the integration of the 1755 cm-1 band in the absence and presence and of 3 M ethanol at a pH of 8.

L02547-9) was used in front of the MCT detector to narrow the IR spectral region and to block scattered light from the 532 nm excitation laser pulse. Spectra in Figures 1 and 2 were collected with 8 cm-1 resolution. To obtain time-resolved data on the µs to ms time scale the DC coupled infrared output of the MCT detector was collected. The data were converted to absorbance difference spectra using a scan of the sample without laser excitation. Figure 1 represents 10 coadditions of the absorbance change of the sample for each interferometric step. The temporal resolution was 5 or 15 µs and collected from -1 ms to 10 ms for some traces and to 100 ms in others. One hundred to 200 time steps were averaged together in each of the time regions representing the pretrigger, 100 µs, 1, 5, and 9 ms. These spectra were extracted and 3-4 trials were averaged together to arrive at the spectra shown in Figure 1. The total signal averaging included 30-40 coadditions and 100-200 time steps. To arrive at Figure 2, the band centered at 1755 cm-1 was integrated in time between 1759 and 1751 cm-1 using the Bruker OPUS data collection software (OS/2 v.3.0.4). Extracted kinetic traces were fit to single or double exponentials using Origin 6.0 software, and the decay times obtained are given in Table 2, for bR with

12908 J. Phys. Chem. B, Vol. 105, No. 51, 2001

Masciangioli and Rice TABLE 2: Assignments of Hydrocarbon Bands in the C-H Stretching Region in BR and DOPhPC with and without Ethanol band frequency (cm-1) spectral assignment CH2 asym str CH3 sym str CH2 sym str CH2 sym str: side chains

bR

BR/EtOH

∆a

DOPhPC

DOPhPC/ EtOH

∆a

2926 2870 undet 2844

2927 2871 undet 2844

+1 +1

2925 2869 2857 2843.5

2926 2870 2858 2843.5

+1 +1 +1 0

0

a ∆ is the wavelength shift due to ethanol rounded to the nearest integer.

Figure 3. Static second derivative spectra showing the comparison of lipid hydrocarbon bands in the C-H stretching region of bR (top) with model lipid DOPhPC (bottom), in the absence (thick lines) and presence (thin lines) of ethanol. Asterisks represent the position of ethanol bands.

Figure 4. Static second derivative spectra showing the comparison of lipid hydrocarbon bands in the methyl and methylene deformation, and wagging region of bR (top) with model lipid DOPhPC (bottom), in the absence (thick lines) and presence (thin lines) of ethanol. Asterisks represent the position of ethanol bands.

TABLE 1: Comparison of Decay Kinetics of Different Spectral Bands for Ethanol Treated and Untreated BR Samples at 25oC band (cm-1)

spectral assignment

pH

BR decay (ms)

BR/EtOH decay (ms)

1755 1755 1524 1185

Asp85 CdO str Asp85 CdO str Retinal trans CdC str Retinal C-C str

7 8 8 8

6.1 ( 0.8 6.8 ( 0.9 6.4 ( 0.6 6.1 ( 0.4

7.0 ( 1.0 18.2 ( 1.6 18.8 ( 4.2 15.3 ( 1.1

and without 3 M ethanol at room temperature. The values given are either the average of the separate runs if the signal-to-noise ratios (S/N) were similar, or the weighted means if the S/N were different. Results Protein Changes: Transient changes of CdO and COOstretching bands of amino acids in bR, associated with M to N and N to O/bR transitions. Bacteriorhodopsin samples with and without 3 M ethanol, at two different pH values, were investigated. Difference absorp-

tion spectra in the CdO stretching region (1800 to 1700 cm-1) after laser excitation of bR are shown in Figure 1. Positive bands in the spectra correspond to features of the newly formed photointermediate(s), and negative bands from the bleaching of ground-state bR molecules. Monitoring the 1760 cm-1 band corresponding to the CdO stretch of the Asp85 residue of bR provides information about proton translocation occurring during the formation and decay of the M and the N intermediates.12 Absorption appears at 1760 cm-1 as M forms. After a few hundred microseconds, the M intermediate transitions to N, causing a shift in the CdO stretching band to 1755 cm-1. In native bR (pH 6-7), the formation and decay of N are ∼1 ms and ∼6 ms, respectively. The kinetics of the M to N transition are determined by monitoring the rise in signal of the N intermediate at 1755 cm-1and at 1398 cm-1. Absorption appears at 1398 cm-1 due to COO- of Asp96.16,27 At pH 7, we do not observe a difference in the N formation above experimental uncertainty in bR with ethanol. At pH 8, with more signal averaging, we observe a difference. Fitting the 1755 cm-1 band, the N intermediate rise (M to N transition) is ∼1000 ( 400 µs without ethanol and ∼600 ( 200 µs with ethanol. Somewhat better data is obtained by fitting the 1398 cm-1 band which yields 1200 ( 40 µs without ethanol and 600 ( 100 µs with ethanol. The result is a 2-fold faster M to N transition in the presence of ethanol. The kinetics of the N to O/bR transition are monitored by observing the rate of decay of the N intermediate at 1755 cm-1. At pH 7, we observe only a 15% longer decay time in bR with ethanol. At pH 8, we observe a significantly longer decay of the N to O/bR transition in ethanol. Comparisons of the results are given in Table 1. Spectral differences and the shift from 1760 to 1755 cm-1 in the Asp85 band at pH 8 are shown in Figure 1. Without ethanol, the decay of the 1755 cm-1 integrated band yields a lifetime of 6.8 ( 0.9 ms. In the presence of ethanol, this value is 18.2 ( 1.6 ms. The result is a 3-fold slower transition from N to O/bR in ethanol. This is shown in Figure 2. The 1398 cm-1 band due to COO- of Asp96 shows similar kinetic behavior, with a decay of 5.3 ( 0.3 ms without ethanol and 18.7 ( 1.5 ms with ethanol. Lower Temperature Measurements associated with the N to O/bR Transition in bR. We also carried out experiments at 10 °C in 3 M ethanol monitoring the 1755 cm-1 band to determine the effect of cooling from room temperature to 10 °C on the N to O/bR transition. Our results indicate a longer decay of 62 ( 14 ms at pH 8. Chromophore Changes: Transient Changes of CdC Stretching of all-trans Retinal Bleach RecoVery in bR. The CdC stretching frequency bleach and recovery at 1524 cm-1, due to the all-trans retinal chromophore is one of the strongest transient bands in the infrared spectrum of bR. This provides a more direct comparison to the visible spectrum kinetics reported by

Dynamics in Bacteriorhodopsin with Ethanol

J. Phys. Chem. B, Vol. 105, No. 51, 2001 12909

TABLE 3: Assignments of Hydrocarbon Bands in the Methylene Scissor and Wagging Region in BR and DOPhPC with and without Ethanol band frequency (cm-1) spectral assignment CH2 def CH3 asym def

Protein band CH2 def (Gly, Asp, Gln, Glu) Lipid band (COO-) sym stretch (Asp85, Asp212, Glu) CH3 sym def, “umbrella” mode CH3 sym def, kink gtg a

bR

BR/EtOH

∆a

1470

1469

-1

1456 1445.5 1437.5 1423

1455.5 1445.5 1437.5 1422.5

-0.5 0 0 -0.5

1393.5

1392.5

-1

1385 1378 1367

1384.5 1378 1366.5

-0.5 0 -0.5

DOPhPC

DOPhPC/EtOH

∆a

undet 1462.5 1453

undet 1463 undet

+0.5

1436.5

1436.5

0

1419

1419

0

1384 1377 1366

1384 1377 1366

0 0 0

∆ is the wavelength shift due to ethanol rounded to the nearest half integer.

Fukuda and Kouyama.23 We observe a bleach and recovery for the bR/buffer/ethanol sample that fits two exponentials slightly better than one with the values 4.1 ( 2.3 and 19 ( 4 ms, respectively. The longer time is similar to the value of the disappearance of the 1755 cm-1 band associated with the N to O/bR transition. This band may have other contributions that could explain the shorter component. The relative amplitudes of the 4 and 19 ms decays are 0.3 and 0.7, respectively. Transient Changes of C-C Stretching at 1185 cm-1 due to Retinal Bleach RecoVery in bR. The transient bleach and recovery at 1185 cm-1 due to a C-C stretching mode in retinal, appears when the Schiff base is reprotonated.17,20,28-30 The bleach corresponds to deprotonation of retinal and the recovery corresponds to reprotonation. The kinetics of recovery of this band should match the Asp96 deprotonation kinetics and correspond to the transition from M to N. The Schiff base of the chromophore is reprotonated from Asp96, but remains in the cis-configuration, in this step. This provides the best comparison to the reported kinetics of the visible spectrum, which is due to the ground-state absorption of the chromophore. We observe times of 6 and 15 ms for the recovery of this band without and with ethanol, respectively. Lipid Structure: Examination and Influence of Ethanol on bR Lipid Vibrational Bands. We observe several shifts of lipid bands in the CH stretching region in the static spectra of bR/ buffer when ethanol is added. The peak positions of these bands based on second derivative spectra are summarized in Table 2 and shown in Figure 3(top). Blue shifts are seen in bands 2870 and 2926 cm-1. These bands are assigned to the CH3 symmetric and CH2 asymmetric stretching frequencies, respectively. Ethanol bands appear at 2983 and 2905 cm-1. The shifts we observe in the bR lipid hydrocarbon bands are small: on the order of 1 cm-1, but are comparable to those seen elsewhere for acyl chain frequencies undergoing gel-liquid crystal phase transitions as a function of temperature.31 To verify the presence of these shifts, we obtain FTIR spectra of the model lipid DOPhPC with and without ethanol. This is an ether-linked branched-chain lipid similar to the predominant native lipids found in bR and allows us to examine the influence of ethanol on one structural lipid in the absence of the bR protein. Second derivative spectra are shown in Figure 3 (bottom). The symmetric CH2, the symmetric CH3, and asymmetric CH2 stretching bands at 2855, 2869, and 2925 cm-1, respectively, are blue shifted with the addition of ethanol. These shifts are similar to those seen when ethanol is added to bR. The region in bR near 1450 cm-1 comprises several CH2 deformation (scissoring) and CH3 asymmetric deformation bands

previously reported at 1466, 1456, and 1442 cm-1.32 We observe bands at 1470, 1456, 1446, and 1438 cm-1. The two bands we report at 1446 and 1438 cm-1 may correspond to the previously reported band centered at 1442 cm-1 or the previously reported band may indeed be two overlapping bands. Our observations are listed in Table 3 and shown in Figure 4 (top). The 1470 and 1456 cm-1 bands in bR are red-shifted upon addition of ethanol. The positions of these bands in the model lipid are different and not easily correlated with the bR bands. Some of the difference may be explained by examining the three bands in the DOPhPC spectrum that are attributed to ethanol alone: 1454, 1420, and 1385 cm-1. These are marked with asterisks in Figure 4 (bottom). The larger intensities of bR bands (Figure 4 top) at 1456 and 1385 cm -1, with or without ethanol, match bands in the DOPhPC-ethanol spectrum (Figure 4 bottom). This suggests that these deformation bands in bR arise from bending vibrations similar to those in ethanol which are present in the protein side chains. Alternatively, they could arise from bending vibrations associated with particular structures found in the DOPhPC lipid-ethanol environment, although we believe this is less likely. We conclude that there are strong bands associated with the bR protein between 1375 and 1475 cm-1. Analyses of the methylene wagging region (1390-1330 cm-1) of the static second derivative FTIR spectra are also summarized in Table 3 and shown in Figure 4. Bands due to isolated (uncoupled) modes in conformationally disordered systems appear in this region. Intensity is generally considered the most sensitive parameter of these bands however, small shifts related to order-disorder do occur.33 Disorder generally results in a shift to higher frequencies. The bands we observe at 1385 and 1367 cm-1 shift to slightly lower frequencies upon addition of alcohol. This could correspond to an ordering of the chains, although this is not in line with expectation. In a more general view, a lower wagging frequency may imply a more hindered motion due to weak protein interaction or hydrogen bonding, solvent interaction, or a structural change. One of these affects may be the origin of the shifts observed here. However, the corresponding bands in the model lipid spectrum do not appear to shift. This suggests the shifts may originate from an ethanol effect on the protein CH2 wagging modes. Alternatively, the shifts may originate from an ethanol effect on minor bR lipid components that are not represented by our model lipid. The observed shifts in this region of the spectrum will need to be substantiated with temperature studies since the shifts are not verified in the model lipid. Another region of interest around 1100 cm-1 where the lipid phosphate headgroup bands appear could not be examined due

12910 J. Phys. Chem. B, Vol. 105, No. 51, 2001 to interference from ethanol absorption bands. Using deuterated solvent also interfered with analyzing ethanol effects on the samples in this region. Previous X-ray lamellar diffraction studies on lipid phosphate headgroups in diphyatanol phosphatidylchholine have been shown to exhibit an order-disorder transition in response to dehydration and temperature changes.34 Discussion The goal of this paper is to correlate changes in lipid or protein structure due to ethanol with changes in the kinetics of bR proton translocation. Examination of the time-resolved vibrational bands in bR allows us to examine the local environments surrounding specific amino acids involved in the proton translocation in addition to the retinal chromophore. We also seek a detailed description of the influence of ethanol on the lipid/bR system. The ethanol effect on the rate of the N to O/bR transition is strong at pH 8; we observe a three times slower rate in the N to O/bR transition of the photocycle of bR with 3 M ethanol added. The time-resolved spectra shown in Figure 1 reveal no noticeable peak shifts indicating no apparent changes in the protein environment between the treated and untreated samples. The lack of a difference in vibrational band peak position with and without ethanol suggests that the protein structure around the Asp85 group is not perturbed by the addition of ethanol. Alternatively, this band may not be sensitive to protein and /or lipid structural changes. The effect of ethanol on the M to N transition is more difficult to observe and is opposite in effect to that seen in the N to O/bR transition. Kaulen35 has noted that the enhancement of the M to N rate is coupled to the slowing of the N to O/bR rate under a variety of perturbations such as mutation or chemical addition. In addition to the ethanol perturbation addressed here and by Fukuda and Kouyama,23 other perturbations include the addition of Triton X-100,36 substitution of Thr46 by Val,37,38 introduction of bulky groups in the helix F,39 and excitation photon dependence. The simplest explanation is that N is stabilized by these additions; however, Kaulen35 attributes this behavior to the stabilization of the “open” N and M forms in comparison with their “closed” forms. We also monitor the changes in the infrared bands associated with the retinal chromophore in addition to the protein bands discussed above. The retinal bands are at 1185 cm-1 (C-C sym stretch in retinal) and 1525 cm-1(CdC sym stretch in all-trans retinal). We observe consistent decays and rises to those seen in the 1755 and 1398 cm-1 bands. Ethanol Effect on Lipid Membrane. One way ethanol may be effecting the dynamics of the bR photocycle may be through altering the interactions of the lipid membrane with the protein. In the analogous rhodopsin photosystem,42,43 Litman and coworkers 3,44 found that alcohols enhanced the formation of the M-like intermediate metarhodopsin II (MII). They attribute this effect to disordering of phospholipid acyl chain packing in the membrane, as well as, disruption of rhodopsin-water hydrogen bonds. Another model of ethanol interaction with biological membranes proposes that hydrogen-bonded water is replaced by ethanol.45,46 Water and ethanol are postulated to compete for hydrogen binding sites on the surface of lipids and proteins, due to similar hydrogen bonding capability of both of them. It is also known that physical properties of the surrounding phospholipid bilayer modulate membrane function. Both lipid conformational order and chain packing can be qualitatively monitored through evaluation of methylene stretch-

Masciangioli and Rice ing frequencies.47 Monitoring shifts in these bands enables one to distinguish between packing and conformational effects. When the bands shift to lower frequency as a result of a change in environment this implies packing geometry alterations, whereas a shift to higher frequencies indicates introduction of gauche rotamers into lipid chains. Our best observation of shifts occurs in the CH2 asymmetric stretch at 2926 cm-1, undergoing a blue shift of 1 cm-1. We also observe a similar shift in the CH3 symmetric stretch at 2870 cm-1. We do not observe any red shifts of these bands, which suggests no alterations to packing of the CH3 and CH2 groups. We do observe blue shifts which suggests changes in the conformation of lipid hydrocarbon chains. The bR sample and model lipid responses to ethanol are relatively consistent in the 3000-2800 cm-1 region (Figure 3). This indicates that the observed changes can be attributed to changes in the lipids. However, this is not the case in the bending (deformation) region of the infrared spectra (Figure 4) where the changes are as likely to be due to interactions with the protein. Here, the shifts in the bR bands do not correspond to model lipid shifts. The shifts in the CH2 deformation mode at 1470 cm-1, where there is not a strong band in the model lipid, the shift in the protein band at 1393.5 cm-1, and possibly the shift in the protein band at 1423 cm-1, suggests the protein is also affected by ethanol. There is further evidence from the literature of the protein being affected by ethanol from absorption anisotropy measurements of retinal in bR membranes. In these studies, ethanol increases the rotational time.26 This is attributed to either increased fluidity of retinal in the protein pocket or increased rotational times of the entire protein in the membrane environment, and arguments are made which support the latter interpretation. Fluorescence anisotropy measurements on 1,6,diphenyl-1,3,5 hexatriene (DPH), a fluorescent molecule (sensor), in the purple membrane in the presence of ethanol, resulted in depolarization parameters that also indicate an increase in fluidity of the protein in the lipid environment in the presence of ethanol. In summary, we show that conformational changes have occurred in the bR lipid due to addition of ethanol. This evidence comes in the form of shifts in the CH3 symmetric and CH2 asymmetric stretching frequencies. We have evidence to suggest the protein may also be effected from the shifts observed in the methylene wagging regions, but this will require follow-up studies. Comparisons with Previous Work. Our native bR sample rates (in the absence of ethanol) for the N to bR transition agree with those reported by Rodig et al.48 Their time constants were determined for the bR photocycle at several pH values in wellhydrated bR films, under similar conditions to ours. At pH 7 and 8 they report N intermediate lifetimes of 5.0 and 5.9 ms, respectively. A pH dependent study of M and N kinetics between pH 3-9 in several alcohols, was previously reported by Fukuda and Kouyama.23 This was carried out by monitoring the chromophore absorption changes in the visible region of the spectrum. We were unable to see the ehtanol effect they reported at pH 6, but by increasing the pH to 8, we observe clear differences in the kinetics due to ethanol. (Higher pH values are known to accentuate the inhibition of the N to O/bR transition.23) Fukuda and Kouyama23 carried out their kinetics at 10 °C, however, we were able to monitor faster kinetics and collected most of our data at room temperature. Fukuda and Kouyama23 found the rate of the M to N transition increased

Dynamics in Bacteriorhodopsin with Ethanol linearly with increasing alcohol concentration, and was accelerated by a factor of ∼3-4 at an alcohol concentration of 3 M. We observed a factor of ∼2 increase with some differences in conditions. They concluded that long-distance proton-transfer involved in the M to N transition (from Asp96 to retinal, ∼10 Å) becomes easier when alcohols are present, due to a softening of the protein conformation by partially breaking hydrogenbonding networks in the protein. We can state by examining the shifting of several lipid vibrational bands that the lipids are affected, however our data suggest these shifts are due to conformational changes in the lipid chains and not necessarily a loosening of the chain packing. Fukuda and Kouyama23 observed a decrease in the rate of N to bR with the addition of alcohols. This decrease was a factor of 3 to 4 times in comparing no ethanol to 3 M ethanol at pH 6 at 10 °C. We observe a decrease of 2.8 times in the N lifetime comparing no ethanol to 3 M ethanol at pH 8 at room temperature. To eliminate the effect of temperature, we examined the N to O/bR decay at 10 °C and report an N lifetime of 62 ms in 3 M ethanol at pH 8. The closest comparison they have is an N lifetime with 2 M ethanol at a PH 8, which appears to be ∼150 ms (2.4 times slower) based on their plotted data. Using their data at 10 °C, bR samples with 3 M ethanol exhibit an N to O/bR decay ∼1.7 times slower than that at 2 M. In conclusion, our N lifetime could be up to four times faster than that observed by Fukuda and Kouyama. 23 Fukuda and Kouyama carried out their experiments in solution and we rehydrated a film several microns thick. We have seen our decays become somewhat longer by increasing the ethanol exposure time, therefore, it is possible that some of the difference may due to the extent of ethanol exposure in our rehydration process. Their samples may be more fully hydrated having been developed in solution. Both experiments were carried out at the same ionic strength of 0.1 M NaCl. Our focus here is to determine mechanistic information, however, this difference may be resolved by determining the two activation energies more precisely. Conclusions We have studied the effect of ethanol on the M to N and N to O/bR transitions at two pH values using time-resolved FTIR spectroscopy. The effect of ethanol on the rate of the M to N transition is an increase in the rate by a factor of ∼2 at pH 8. The effect on the rate of the N to O/bR transition in the proton pump sequence is a decrease in the rate by a factor of 3. The absolute decay rates differ from previously reported data from visible spectroscopy of the retinal absorption changes, however, the trends hold up and are substantiated on a molecular level using infrared spectroscopy. Correlating the kinetic changes to protein and lipid spectral changes due to the addition of ethanol proves more challenging. Previous reports mention that alcohols “soften” the lipids.23 We present direct evidence that the lipids in bR are affected by the addition of ethanol. We interpret this as a conformational change and not a loosening of the chain packing. This may occur because the lipids in bR are already less tightly packed due to the methyl groups. Ethanol clearly affects the bR proton pumping dynamics, however, it remains to be determined whether this is predominantly through changes in the lipids or changes in the protein. On the basis of the kinetics affected by the addition of ethanol, and the data in hand, the most likely place for direct protein influence in on the CP side of the protein. The current literature does not provide any evidence that ethanol molecules penetrate

J. Phys. Chem. B, Vol. 105, No. 51, 2001 12911 into the CP side of the protein and affect hydrogen bonding or proton transport in a direct way. It is a distance of about 6 Å from Asp96, the proton donor to the retinal chromophore. We observe changes in the bR lipids due to ethanol, and we observe changes in bR not associated with DOPhPC-type lipids. We see a large effect on the proton transport rates on the CP side of bR. However, we cannot assign cause/effect to these two observations. We can say, however, that the band positions of Asp85 on the EC side, Asp96 on the CP side, the retinal C-C symmetric stretch, and the retinal CdC symmetric stretch, which monitor the M to N and N to O/bR transitions, are not noticeably shifted. This suggests that the protein in these regions (the retinal pocket, slightly toward the EC side where Asp85 is located, and the CP side of protein near Asp96) has not undergone any noticeable structural changes affecting these two transitions. Acknowledgment. We gratefully acknowledge Professor Mostafa El-Sayed and Dr. Jianping Wang at Georgia Tech for supplying us with purple membrane samples. This work was performed while T.M.M. held a National Research CouncilNaval Research Laboratory Research Associateship. We also thank the Office of Naval Research for funding this work through the Naval Research Laboratory. References and Notes (1) Gutman, M.; Nachliel, E. Annu. ReV. Phys. Chem. 1997, 48, 32956. (2) Bourne, H. R.; Meng, E. C. Science 2000, 289, 733-734. (3) Mitchell, D. C.; Litman, B. J. J. Biol. Chem. 2000, 275, 53555360. (4) Oesterhelt, D.; Stoeckenius, W. Nature New Biol. (London) 1971, 233, 149-52. (5) Stoeckenius, W.; Bogomolni, R. A. Annu. ReV. Biochem 1982, 51, 587-616. (6) Lozier, R. H.; Bogomolni, R. A.; Stoeckenius, W. Biophys. J. 1975, 15, 955-62. (7) Kates, M. In Biological Membranes: Aberrations in Membrane Structure & Function; Karnovsky, M. L., Leaf, A., Bolis, L. C., Eds.; Alan R. Liss, Inc.: New York, 1988; pp 357-384. (8) Kamekura, M.; Kates, M. In Halophilic Bacteria; Rodriguez-Valera, F., Ed.; CRC Press: Boca Raton, 1988; Vol. II.; pp 25-54. (9) Kates, M. Experiencia 1993, 49, 1027-1036. (10) Lindsey, H.; Petersen, N. O.; Chan, S. I. Biochimica et Biophysica Acta 1979, 555, 147-167. (11) Joshi, M. K.; Dracheva, S.; Mukhopadhyay, A. K.; Bose, S.; Hendler, R. W. Biochemistry 1998, 37, 14 463-14 470. (12) Braiman, M. S.; Mogi, T.; Marti, T.; Stern, L. J.; Khorana, H. G.; Rothschild, K. J. Biochemistry 1988, 27, 8516-20. (13) Fahmy, K.; Weidlich, O.; Engelhard, M.; Tittor, J.; Oesterhelt, D.; Siebert, F. Photochem. Photobiol. 1992, 56, 1073-83. (14) Metz, G.; Siebert, F.; Engelhard, M. FEBS Lett. 1992, 303, 23741. (15) Gerwert, K.; Hess, B.; Soppa, J.; Oesterhelt, D. Proc. Natl. Acad. Sci. U.S.A. 1989, 86, 4943-7. (16) Bousche, O.; Braiman, M.; He, Y. W.; Marti, T.; Khorana, H. G.; Rothschild, K. J. J. Biol. Chem. 1991, 266, 11 063-7. (17) Fodor, S. P. A.; Ames, J. B.; Gebhard, R.; Van den Berg, E. M. M.; Stoeckenius, W.; Lugtenburg, J.; Mathies, R. A. Biochemistry 1988, 27, 7097-101. (18) Otto, H.; Marti, T.; Holz, M.; Mogi, T.; Lindau, M.; Khorana, H. G.; Heyn, M. P. Proc. Natl. Acad. Sci. U.S.A. 1989, 86, 9228-32. (19) Ames, J. B.; Mathies, R. A. Biochemistry 1990, 29, 7181-90. (20) Pfefferle, J. M.; Maeda, A.; Sasaki, J.; Yoshizawa, T. Biochemistry 1991, 30, 6548-56. (21) Sasaki, J.; Lanyi, J. K.; Needleman, R.; Yoshizawa, T.; Maeda, A. Biochemistry 1994, 33, 3178-84. (22) Tavan, P.; Schulten, K.; Oesterhelt, D. Biophys. J. 1985, 47, 41530. (23) Fukuda, K.; Kouyama, T. Photochem. Photobiol. 1992, 56, 105762. (24) Chiou, J.-S.; Krishna, P. R.; Kamaya, H.; Ueda, I. Biochim. Biophys. Acta 1992, 1110, 225-233. (25) Mitaku, S.; Ikuta, K.; Itoh, H.; Kataoka, R.; Naka, M.; Yamada, M.; Suwa, M. Biophys. Chem. 1988, 30, 69-79. (26) Kikukawa, T.; Araiso, T.; Shimozawa, T.; Mukasa, K.; Kamo, N. Biophysical Journal 1997, 73, 357-366.

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