Interactions of Hydration Water and Biological Membranes Studied by

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J. Phys. Chem. B 1999, 103, 8036-8050

FEATURE ARTICLE Interactions of Hydration Water and Biological Membranes Studied by Neutron Scattering J. Fitter,*,†,‡,| R. E. Lechner,§ and N. A. Dencher*,†,⊥ Institut fu¨ r Biochemie, Technische UniVersita¨ t, Petersenstr. 22, D-64287 Darmstadt, Germany; Forschungszentrum Ju¨ lich, IBI-2, Biologische Strukturforschung, D-52425 Ju¨ lich, Germany; Hahn-Meitner Institut, BENSC, Glienicker Str. 100, D-14109 Berlin, Germany ReceiVed: April 14, 1999; In Final Form: June 30, 1999

To elucidate general features of structural and dynamical properties of hydration water and the influence of hydration water on the dynamical behavior of biomembranes, purple membranes from halobacteria and disk membranes from bovine retinae have been studied by neutron scattering techniques. Hydrated films of oriented multilamellar membrane stacks were used to measure lamellar diffraction patterns and quasielastic incoherent neutron scattering as a function of hydration level, of temperature, and of the protein/lipid ratio. These measurements revealed a strong interaction of a “first hydration layer” with the membrane surface and a reduced self-diffusion of aqueous solvent parallel to the membrane surface (the self-diffusion coefficient is about 5 times smaller as compared to excess water). The picosecond internal molecular motions of the protein/ lipid complex are strongly affected by the amount of solvent interacting with the lipids and the membrane proteins. In particular, the lipids and their ability to attract solvent molecules play an important role for “hydration-induced flexibility” of biomembranes. On the basis of these measurements, the impact of the hydration process on the function of biomembranes is discussed for the light-driven proton pump bacteriorhodopsin in purple membranes.

1. Introduction Biological membranes are complex and heterogeneous proteinlipid assemblies separating living cells from their extracellular surroundings, i.e., these membranes represent the interface between the cell and the environment. In addition, all eukaryotic cells (cells of plants and animals) contain a number of compartments (organelles) such as the cell nucleus, mitochondria, or chloroplasts, etc., which are also bounded by membranes. The basic function of biomembranes is to provide different spatial compartments and to act as highly selective barriers for the exchange of ions and molecules between the different compartments. Because of their diverse biological functions, such as energy conversion, signal transduction, or material transport, etc., biomembranes are composed of individual mixtures of many lipids and of specific amphiphilic proteins. The basic structural element is provided by a lipid bilayer of 40-50 Å thickness (see Figure 1) in which the proteins are embedded (membrane proteins) or to which the proteins are attached (see for example ref 1). The aqueous solvent environment provides not only an entropic driving force (hydrophobic interaction) for the structural self-assembly process of lipid bilayers, but also has profound influences on the * Corresponding author. | Phone: +49-2461-612036. Fax: +49-2461-612020. E-mail: j.fitter@ fz-juelich.de. ⊥ Phone: +49-6151-165275. Fax: +49-6151-164171. E-mail: dencher@ pop.tu-darmstadt.de. † Institut fu ¨ r Biochemie, Technische Universita¨t. ‡ Forschungszentrum Ju ¨ lich, IBI-2, Biologische Strukturforschung. § Hahn-Meitner Institut, BENSC.

dynamics and thermodynamics of such systems. (Analogous to lipid bilayers, aqueous solvent and hydrophopic interaction play also a major role for the polypeptide chain folding in globular proteins and for the conformational stability and flexibility of proteins.) Beside the hydrophobic interaction which initiates the bilayer formation, van der Waals interactions between the apolar hydrocarbon-chains and electrostatic interactions as well as H-bonds between water molecules and polar headgroups stabilize the cooperative structure of lipid bilayers. The influence of the solvent environment on structural, dynamical, and functional properties of biomolecular systems (i.e., biological membranes, proteins, nucleic acids, etc.) is evident from numerous experiments performed on many systems as a function of the degree of hydration [see, for example, refs 2-4]. Although we will discuss biological membranes here, many characteristics of hydration and hydration water were also observed with globular proteins and seem to be general properties of all biological systems.4 In general, there are two main areas of studies dealing with questions concerning characteristics of the hydration process: (i) properties of aqueous solvent in the vicinity of biomolecules (e.g., structural and dynamical properties of hydration water), and (ii) the influence of the solvent on the structure, dynamics, and function of biomolecules (e.g., dependence on hydration level). Both aspects have been studied by the use of various techniques, such as NMR spectroscopy,2,5-7 dielectric relaxation,8 neutron and high-resolution X-ray diffraction,9-11 molecular dynamic simulations,12-16 calorimetric measurements,17,18 infrared spectroscopy,19 and neutron spectroscopy.20-24 On the basis of NMR data, Kuntz and co-workers introduced already

10.1021/jp9912410 CCC: $18.00 © 1999 American Chemical Society Published on Web 09/03/1999

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J. Phys. Chem. B, Vol. 103, No. 38, 1999 8037

Figure 1. Schematic representation of a multilamellar stack of purple membrane (PM) sheets (left) and schematic clipping enlargements of hydrated purple membranes (right) top: side view of bacteriorhodopsin (BR) molecules embedded in lipids (without exact relative scaling) forming the purple membrane. The inter-bilayer space is filled with water molecules. The one-dimensional lamellar periodicity dl includes the thickness of the membrane and the “thickness” of the inter-bilayer water. Depending on the hydration level, dl varies from 49.5 Å to more than 100 Å. (right) bottom: top view on an electron density map of a projected structure of three BR molecules (trimer) in the two-dimensional hexagonal lattice cell. The narrow iso-lines (high electron density) represent a projection of seven helices as part of each BR molecule. Purple membranes hydrated at 98% r.h. exhibit an in-plane cell parameter ds ) 62.5 Å.

in the early seventies a crude classification of water molecules characterized by different interactions with the biomolecule:2 (I) water which is qualitatively similar to excess water (bulk water) with respect to translational and rotational freedom (moderate or weak interaction with the biomolecule), (II) “bound water” which is translationally restricted to the surface of the biomolecules (“hydration shell”) or to larger cavities (e.g., channels) inside the biomolecules, and (III) “nonrotational bound water” which is essentially “site bound” to the biological macromolecule via more than one H-bond. All these categories of water molecules have been studied using very different approaches and techniques ranging from localization of single water molecules inside a protein structure (category III)9,11 to analyzing dynamical and thermodynamical properties of hydration water (categories I and II) in the framework of glass transition17-19 and mode coupling theory. Although so many studies have already been performed on aspects of the hydration process in the past, there is still intensive activity in this field. In particular, the role of the different categories of water with their specific properties and the influence of these water molecules on the biomolecule for the function of the biomolecule is the focus of many investigations.4,25-30 Results concerning various aspects of the hydration process as obtained from neutron scattering experiments are described in the present article. The intention of the present article is to

outline the impact of neutron scattering studies for understanding fundamentals of the hydration process of biomembranes. Incoherent neutron scattering which provides information on the entity of all water molecules located in the sample (and to a large extent interacting with biomembranes) is a complementary method to other powerful methods, such as X-ray diffraction, NMR spectroscopy, or FTIR, studying individual (more or less bound) water molecules (see, for example, refs 4-7,911). 2. Neutron Scattering Techniques to Study Biological Membranes 2.1. Purple Membranes and Disk Membranes. Bacteriorhodopsin (BR), which is the prototype of an integral membrane protein, is the only protein in the so-called purple membrane (PM) of Halobacterium salinarum and forms, together with few lipids, highly ordered two-dimensional lattices (see Figure 1). It functions as a light-driven proton pump and is one of the best-characterized membrane proteins. Halobacteria use bacteriorhodopsin to generate a proton electrochemical potential across the cell membrane which is utilized by the organisms as a driving force for ATP synthesis.31 By high-resolution electron crystallography on natural two-dimensional purple membrane crystals32,33 and by X-ray diffraction on three-dimensional

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TABLE 1: Distribution of Protons in the Purple Membrane from Halobacterium salinarum and in the Disk Membrane from Bovine Rod Outer Segmentsa purple membrane

composition BR Rh lipids other proteins total

disk membrane

number of number of content nonexchangeable content nonexchangeable (w/w) protons (per (w/w) protons (per [%] BR-molecule) [%] RH-molecule) 75

1891

25

635

100

2526

45 50 5 100

2619 3621 131 6371

a

The numbers were calculated using the knowledge of bacteriorhodopsin (BR) and rhodopsin (RH) amino acid compositions and of the protein/lipid compositions in both membranes.48-51

crystals,34-36 the 3D structure of bacteriorhodopsin has been determined to a resolution of 2.3 Å. Bacteriorhodopsin consists of a single polypeptide chain of 248 amino acids (molecular mass of 26.5 kDa) traversing the lipid bilayer (only 10 lipids per protein) in seven helical segments.37 The application of many biophysical methods demonstrated that bacteriorhodopsin in the purple membrane was, and still is, an appropriate biomolecular system for extending our knowledge of various relevant topics, such as (i) structure-function relation of proteins,38-42 and (ii) the role of hydration water in bioenergetics.43-47 The photoreceptor rhodopsin (RH) is located in the disk membrane (DM) of retinal rod photoreceptor cells (see, for example, ref 1). Similar to bacteriorhodopsin, rhodopsin is an integral membrane protein having a seven-helix membranespanning structure but with a larger molecular mass of about 40 kDa. Although rhodopsin and bacteriorhodopsin, which show many similarities, are the only or the dominant fractions of membrane proteins in their specific membranes (bacteriorhodopsin: 100% (w/w) in the purple membrane and rhodopsin: 95% (w/w) in the disk membrane), the disk membranes and the purple membranes are rather different. Disk membranes are characterized by a much “larger fluidity” due to more lipid molecules per rhodopsin molecule (≈60 mol lipids/mol rhodopsin) as compared to the purple membrane, where immobilized trimes of bacteriorhodopsin form a rigid protein-lipid complex (see also Table 1). Further, the membrane surface of the disk membrane is rather different as compared to the purple membrane, because rhodopsin has larger loop regions (part of the polypeptide chain which connect the helix regions) protruding into the inter-membrane space. More details concerning the preparation of both types of membrane sheets (purple membranes from halobacteria and disk membranes from bovine retinae) from the organisms are described elsewhere.52,53 With respect to neutron scattering experiments, it is essential that both types of membranes need to be available in amounts of a few hundred milligrams. A sample of membranes can be obtained as hydrated multilamellar stacks with membranes parallel to the surface of a sample support (Figure 1). Theses samples are made of alternating layers of membrane sheets and of water molecules. With respect to our purpose, these samples reveal several advantages. (i) Lamellar membrane stacks, even if the sample is hydrated at low levels (down to only two layers of water between two adjacent membrane sheets), represent a sample which shares important properties with “natural membranes” (i.e., membranes in solutions), e.g., in many cases fully functional membrane proteins, no artificial protein-protein contacts (often occurring in hydrated powders of globular proteins). (ii) Lamellar membrane stacks, also at different hydration levels, can be used to study

the function of bacteriorhodopsin and of rhodopsin (for example by time-resolved absorption spectroscopy54) and structural as well as dynamical properties of the lipids, of the membrane proteins, and of the aqueous solvent molecules. (iii) Lamellar membrane stacks represent a partly oriented sample. Therefore one can study many properties as a function of orientation (e.g., diffusion of solvent molecules parallel and perpendicular to the membrane surface, see Section 3.1). All neutron scattering experiments, which are described in the following sections, have been performed with these kind of hydrated multilamellar membrane sheets. 2.2. Neutron Diffraction with Lamellar Membrane Stacks. For investigating properties of hydration water and interactions of hydration water with biological membranes, it is of particular interest to determine the thickness of the inter-membrane space (usually filled with solvent) as a function of the hydration level and as a function of temperature. To measure this quantity, lamellar diffraction patterns were recorded by the use of a neutron diffractometer in so-called ω-2θ scans (for more details see, for example, ref 55). The one-dimensional periodicity dl (see Figure 1) along the direction perpendicular to the membrane surfaces causes lamellar reflections which were recorded with a two-dimensional position-sensitive 3He-detector. The samples, hydrated with D2O, exhibit sufficient intensities for first- and second-order reflections after a measuring time of a few minutes. Samples were hydrated with D2O instead of H2O, because measurements with D2O samples exhibit less incoherent background scattering as compared to H2O samples. Values of the lamellar spacing dl were obtained from 2θ values using Bragg’s law:

nλ ) 2dl sin θ

(1)

where λ is the wavelength of the incoming neutrons, 2θ is the angle between incoming and scattered neutrons, and n is the order of reflections. Temperature (T)- and hydration (h)dependent values of the lamellar spacings revealed important properties of the aqueous solvent as demonstrated in Section 3.2. In the case of purple membranes, where bacteriorhodopsin forms a natural two-dimensional hexagonal lattice, neutron diffraction measurements provide also structural information about bacteriorhodopsin and the lipids (see Figure 1). The distinct intensity distribution at reciprocal lattice points (indexed in the h,k-plane) corresponds to the structure and the arrangement of BR and the lipids in the “in-plane” unit cell (cell dimension ds, see Figure 1) in real space.56,57 The application of H/D contrast variation (i.e., taking advantage of the large difference of coherent scattering length between H with bcoh ) -0.37 × 10-12 cm and D with bcoh ) +0.667 × 10-12 cm) using samples hydrated with H2O and D2O, enables one to determine the amount and the localization of water bound to bacteriorhodopsin or to the lipid headgroups.46,47 2.3. Incoherent Neutron Scattering. With respect to studies on biological membranes, neutrons reveal a very important property. They combine a wavelength of the order of atomic or molecular dimensions (1-10 Å) with a low energy (300-0.8 meV), which makes it possible to study both the structure and dynamics of a system simultaneously (see, for example, ref 58). While diffraction experiments focus on the elastic coherent part of the scattering process (Section 2.2), the use of neutron spectrometers provides information about the dynamics due to inelastic and quasielastic scattering. Neutron spectroscopy using time-of-flight (tof) and backscattering spectrometers permits the investigation of motions in the time range from 10-14 to 10-9

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s. On this time-scale the spectrum is dominated by various types of thermal fluctuations from small-amplitude atomic and molecular vibrations up to large-amplitude stochastic reorientational motions of molecular subunits (e.g., polypeptide side groups).58-62 Because of the large incoherent cross section of hydrogen nuclei (i.e., σinc(H) ) 79.9 [barns] compared to, for example, σinc(D) ) 2.04, σinc(C) ) 0.001, σinc(N) ) 0.49) and the fact that hydrogen atoms are distributed “quasi-homogeneously” in the biological macromolecule, this technique is a powerful tool for the study of all internal motions. In the case of hydrated membranes, the technique enables us to study two aspects of interest separately: (i) internal molecular motions of the biological macromolecules (i.e., proteins and lipids), and (ii) motions of the solvent molecules (H2O). In the former case, the samples need to be hydrated with D2O. Then the predominant part of the measured signal is incoherent neutron scattering (INS) due to nonexchangeable hydrogen atoms. [Note, that the relevant total scattering cross section of H (i.e., incoherent plus coherent scattering cross section) is about eleven times larger than that of D.] Many of the charged and polar polypeptide side groups as well as the polar lipid headgroups contain exchangeable protons, which are replaced by deuterons. Nevertheless, most of the protons are not exchangeable, and from a practical point of view, what is relevant for the incoherent neutron scattering, these protons are distributed nearly homogeneously in the sample (see Table 1). In contrast to smallangle neutron scattering (SANS) techniques and other neutron diffraction methods (Section 2.2), in INS experiments we are not dealing with H/D contrast matching, because coherent scattering effects, such as Bragg reflections and small angle scattering, are excluded from the analyzed data. Here we have to consider the individual incoherent scattering contributions of all hydrogen atoms (which are summed over all H), and which therefore “monitor” the general dynamical properties of the sample as a probe (more methodical details see, for example, ref 59). The dynamical behavior of the solvent molecules can be studied by measuring two samples, one hydrated with D2O, and the other hydrated with H2O. A subtraction of spectra measured with D2O from those measured with H2O gives to a first approximation data, which mainly represent scattering from the solvent H2O (see Section 3.1). In all these measurements the double-differential cross section,

δ2σ 1 |k1| 2 ) ‚[b ‚S (Q,ω)] δΩδω 4π |k0| inc inc

(2)

determines the number of neutrons scattered into a solid angle element δΩ and an energy transfer element δω. Here binc is the incoherent scattering length, while k0 and k1 are the wave vectors for the incident and scattered neutrons, respectively (with momentum transfer Q ) k1 - k0). Information on the dynamics of individual hydrogen atoms can be obtained from the incoherent scattering function Sinc(Q,ω) using the formalism of self-correlation functions developed by Van Hove.63 The selfcorrelation function Gs(r,t) is the Fourier transform in space and in time of the incoherent scattering function,

Sinc(Q,ω) )

∫-∞∞e-iωt∫-∞∞eiQr‚Gs(r,t)drdt

1 2π

(3)

In the classical approximation (see, for example, refs 58,60) Gs(r,t) describes the average time-dependent probability density distribution of N hydrogen atoms,

N

〈δ[r + Ri(0) - Ri(t)]〉 ∑ i)1

Gs(r,t) ) N-1‚

(4)

where Ri(0) and Ri(t) are position vectors of atom i for the time t ) 0 and for the time t. In general, it is not possible to calculate the trajectories Ri(t) directly from the measured scattering function, so that in practice special analytic models are used to investigate the movements of the individual atoms. Using this kind of analysis, a calculated theoretical scattering function Stheor(Q,ω) is fitted to a measured scattering function Smeas(Q,ω),

Smeas(Q,ω) ) F‚e-pω/2kBT‚[Stheor(Q,ω) X Sres(Q,ω)] (5) where a convolution (X: energy convolution operator) with the resolution function Sres(Q,ω) (obtained from measurements of vanadium as a standard for “pure elastic incoherent” scattering) and factors, such as the normalization factor F and the detailed balance factor exp(-pω/2kBT) have been applied. According to the stochastic character of the predominant part of the motions (with samples at physiological conditions), the following expression is used for the theoretical incoherent scattering for localized diffusive motions (case (i), internal motions of proteins or lipids), i.e., when long-range diffusion is absent:

Stheor(Q,ω) ) e-〈u 〉Q ‚[A0(Q)‚δ(ω) + 2

2

∑n An(Q)‚Ln(Hn,ω)]

(6)

Here the scattered intensity is separated into an elastic δ(ω)shaped component (experimentally observed with the resolution width Γres) and quasielastic Lorentzian-shaped contributions Ln(Hn,ω), parametrized by the width Hn ) (τn)-1 (τn are the corresponding correlation times) and the quasielastic incoherent structure factors An. The intensity of the elastic component is given by the elastic incoherent structure factor (EISF) A0, while the An give the quasielastic intensities (QISF). Faster motions are taken into account by the Debye-Waller factor, exp(-〈u2〉‚ Q2), where 〈u2〉 gives the global average “mean square displacements” of vibrational motions of all H-atoms. Instead of a phenomenological interpretation of A0, An, and Hn, theses values can also be interpreted in terms of specific models, applied to describe the specific types of motion (see, for example, Section 4.3). In the presence of long-range translational diffusion (TD), as part of the solvent movements [case (ii)], a convolution of the above-mentioned scattering function for localized motions with the TD-scattering function, STD(Q,ω), is required; this gives

Stheor(Q,ω) ) e-〈u 〉Q ‚[A0(Q)‚STD(Q,ω) + 2

2

∑n An(Q)‚Ln(Hn,ω) X STD(Q,ω)]

(7)

Scattering functions for translational diffusion are often derived from the Chudley-Elliott (CE) jump-diffusion model. In its simplest form this theory starts from the description of a random walk of the diffusing particle on a Bravais lattice of sites available for diffusion,64 assuming equivalence of all sites, negligible jump time, and absence of correlations between different diffusing particles and between successive jumps. The corresponding formula for single crystals is

STD(Q,ω) )

HTD(Q) 1 π |H (Q)|2 + ω2 TD

(8)


Fitter et al.

where the width of this Lorentzian function is given by

HTD(Q) )

11 τn

n

[1 - exp(-iQdj)] ∑ j)1

‚

(9)

dj are the n jump vectors joining a site with its n nearest neighbor sites and τ is the mean residence time per site of the diffusing particle. At small scattering angles, STD(Q,ω) behaves according to translational diffusion in a continuum, since

limQf0 HTD(Q) ) Ds‚Q2

(10)

that is,

STD(Q,ω) )

Ds‚Q2 1 π [D ‚Q2]2 + ω2

(11)

s

in the low-Q regime. For disordered systems, such as polycrystalline powders, liquids, etc., isotropic approximations of eq 8 can be used where, in the case of three-dimensional diffusion, eq 9 is replaced by

HTD(Q) )

[

]

sin(Qd) 1 1τ Qd

(12)

with d ) jump-distance and 1/τ ) 6Ds/d2; Ds ) self-diffusion coefficient in three dimensions. In the case of isotropic two-dimensional diffusion in a plane, such as the surface of a biological membrane (see Section 3.1), the orientation of this plane with respect to the scattering vector Q has to be taken into account and the corresponding expression is then

HTD(Q) )

[

]

sin(Qpd) 1 1τ Qpd

(13)

where Qp are the components parallel to the membrane plane of the scattering vector Q and 1/τ ) 4Ds/d2; Ds ) self-diffusion coefficient in two dimensions. 3. Structural and Dynamical Properties of Aqueous Solvent 3.1. Diffusion of Inter-Membrane Water. In stacks of hydrated biomembranes a certain amount of water is located inside the membrane proteins or attached to lipid between two adjacent headgroups.46,47,65,66 In addition to this intra-membrane water, another category of generally more “bulky” intermembrane water is found between two adjacent bilayers. The dynamical properties of the inter-membrane water were studied with purple membranes hydrated at a level which (i) should give a scattering signal due to solvent scattering, that is strong enough to be analyzed in difference spectra (PM/H2O-PM/ D2O) (g10% of the total PM/H2O scattering), and (ii) provides a study of mainly that water component which is interacting with the membrane.23,67 If the hydration level is very high, too many water molecules behave like excess water, which makes it impossible to study molecules belonging to the “hydration water”. (By analyzing difference spectra, we are in general only able to study the “total scattering” representing dynamical parameters, which are the average over all solvent molecules.) A proper hydration level for our purpose was achieved with samples equilibrated at 98% relative humidity (r.h.) which corresponds h ≈ 0.35 g of H2O per g of purple membrane (equivalent to approximately 650 water molecules per bacterio-

rhodopsin molecule, where bacteriorhodopsin is completely functional). At this hydration level only a minor amount of water is located in the interior of bacteriorhodopsin, while the major amount of water is found on the membrane surface and in the inter-bilayer space (with dl ) 61.5 Å and a thickness of approximately 49.6 Å for the sole membrane the water layer thickness is approximately 12 Å, see Figure 1). Both samples, H2O- and D2O-hydrated, were measured at room temperature (T ) 293 K) at the inverted time-of-flight spectrometer IRIS (ISIS-facility, Didcot, GB). To analyze a potential anisotropy of the water diffusion, measurements with two different sample orientation angles R ) 45° and R )135° have been performed. The resulting difference spectra for a scattering angle φ ) 90° (corresponding to the angle between k0 and k1) are shown in Figure 2. This figure shows the quasielastic scattering due to water molecules fitted by a theoretical scattering function including local rotational diffusion and translational diffusion of water molecules (see eqs 7, 8, and 13). The advantage of φ ) 90° with respect to the chosen sample orientations is, that for R ) 45° the momentum transfer Q is perpendicular to the membrane plane and for R ) 135° Q is parallel to the membrane plane (see insets of Figure 2). In addition to neutron scattering data, results from pulsed-field gradient (PFG) NMR have been included to determine a more precise Ds-value.23 In combination with these Ds-values, the spectra were fitted using the equations given in Section 2.3. In Figure 3 the values of the line width HTD are plotted as a function of Q2 for both sample orientations, together with theoretical curves according to eq 13. The analysis revealed the following results. (i) Long-range translational diffusion of water molecules is anisotropic and occurs only parallel to the membrane plane within the limit of the given time window (1-50 ps). The data are well-fitted by assuming that the major part of the interbilayer water (95%) participates in diffusion parallel to the membrane surface with a self-diffusion coefficient Ds ) 4.4 × 10-6 cm2/s ((3%) which was determined by PFG NMR and matches rather well the HTD values in the low Q-range (Figure 3). This diffusion is slowed by a factor of 5 as compared to diffusion in (pure) excess water. The translational jump distance d was found to be about 4.1 Å, i.e., three times larger than that of excess water68 at the same temperature. Such a large value is plausible, if this average nearest-neighbor distance corresponds to a distance between potential minima relevant for the diffusive motion on this time scale (i.e., binding sites for water molecules on the membrane surface). (ii) The translational diffusion is accompanied by a local diffusive proton motion, presumably related to the rotation of water molecules. This local rotational motion was found to be about six times slower than the corresponding type of motion in (pure) excess water.68 Qualitatively, the measured data further imply that rotational motions parallel to the membrane plane (i.e., rotation axis perpendicular to the membrane surface) have a higher rate than motions perpendicular to the membrane plane (for more details see refs 23,67). In summary, we found that the inter-bilayer water is clearly affected by interactions with the membrane surface. According to the crude classification of water in the vicinity of biomolecules given in the Introduction, the major part of the interbilayer water in purple membranes shares some similarities with that of category I. Although the inter-membrane water is characterized by a decreased diffusion coefficient and decreased rates of local rotational motions as compared to excess water, the major part of this water still performs translational and rotational motions within the given time window. A comparison

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Figure 3. Translational diffusion line widths HTD plotted as a function of Q2 . The experimental values are given for sample orientation angle R ) 135° with Q parallel to the membrane surface (solid circles) and for R ) 45° with Q perpendicular to the surface (open squares). The points are rather well approximated by the theoretical HTD (solid lines) calculated by using eq 12. The dotted line approximates the “continuous diffusion model” for R ) 135°, valid only in the low-Q limit.23

Figure 2. Quasielastic scattering related to translational and rotational diffusion of water molecules. This figure shows a fit of a “differencespectrum” measured with scattering angle φ ) 90.4° and sample orientation of (a) R ) 45°, i.e., Q perpendicular to the membrane surface and of (b) R ) 135°, i.e., Q parallel to the membrane surface (see insets). The spectra (experimental points: open circles) were obtained by subtraction of a spectrum measured with purple membranes in D2O from a spectrum of purple membranes measured in H2O (both samples hydrated at h ) 0.35). The area between the upper solid line (fitting the experimental points) and the dotted line represents the elastic scattering broadened by translational diffusion (TD) (Lorentzian LTD with line width HTD given in Figure 3). The quasielastic scattering due to rotational motions (L1 with line width H1, see eq 7) is represented by the area between the dotted line and the dashed line. H1, which corresponds to the rotational motion, is approximately 45 µeV for R ) 45° and about 65 µeV in the case of R ) 135°. A linear background is represented by the area between the dashed line and the horizontal axis. Although the spectra are not fitted perfectly (difference spectra exhibit larger statistical errors), a comparison of both spectra clearly reveals a broader line width HTD for R ) 135° as compared to R ) 45°.23

of water on the surface of other membranes or of different biomolecules (e.g., globular proteins), also studied with various techniques and at partly different hydration levels, is crucial; nevertheless, it is interesting to note that several studies revealed similar values for self-diffusion coefficients ranging from Ds

) 10-6 -10-5 cm2/s.8,15,21,69,70 Beside a general relevance of solvent properties for all types of biomolecules, the water mobility is of particular interest in the case of the purple membrane, because bacteriorhodopsin is a proton pump and water molecules might play an important role as charge carriers (OH-, H3O+) for bioenergetic processes. The possible impact of water mobility on proton migration along the membrane surface will be discussed in more detail in Section 5. 3.2. Temperature-Induced Dehydration-Rehydration Transition. To analyze the solvent-membrane interaction, temperature-dependent features of the hydration water, such as supercooling or solvent freezing, were investigated. For this purpose the lamellar spacing dl of purple membrane and disk membrane multilayer systems has been measured with neutron diffraction as a function of temperature. A more detailed and systematic study has been performed with purple membranes, where samples with four different levels of hydration were measured.71 Figure 4 shows the temperature dependence of dl as measured for samples which had been equilibrated at room temperature at the following different relative humidities (D2O vapor): 100% (h ≈ 0.5), 94% (h ≈ 0.3), 86% (h ≈ 0.2), and 0% (h ≈ 0.06). During the cooling cycle (open symbols in Figure 4) of samples equilibrated at 100% r.h., starting at 295 K, dl stays approximately constant down to a few degrees below the freezing point of bulk water (Tf ) 273.15 K for pure H2O and 276.97 K for pure D2O). Then, at a temperature denoted by Tfh, a discontinuity occurs, which is connected with a large decrease in dl by a step of the order of 20% to 30% (Figure 4). Below Tfh, further cooling is accompanied by a much weaker continuous decrease of dl. The slope of this decrease diminishes upon cooling and becomes very small below about 240 K. Upon reheating the sample, dl follows a continuous curve, starts to increase noticeably near a temperature Tmh* ≈ 255 K, and exhibits a hysteresis-type behavior above this temperature. It does not show a step at the temperature Tfh, but a discontinuity now occurs at a somewhat higher temperature denoted by Tmh (but still below the melting point of bulk D2O). At Tmh the lamellar lattice constant reaches a value close to the level


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Figure 4. Lamellar lattice constant dl as a function of temperature, observed in cooling/heating cycles for four different purple membrane samples, that had been equilibrated at room temperature with different relative humidities (D2O): 100% r.h., 94% r.h., 86% r.h., and 0% r.h. The freezing and melting discontinuities temperatures, Tfh and Tmh, of the membrane water are indicated, using the same symbols for 100% r.h. and for 94% r.h.; Tmh* is the approximate starting point of the membrane rehydration region. Tf denotes the freezing temperature of pure D2O at 276.97 K. Open symbols: cooling, full symbols: heating branches of the measurements; lines have been drawn through the experimental points to guide the eye.71

originally observed at room temperature and stays constant beyond Tmh. This behavior is perfectly reproducible. For samples hydrated at r.h. e94%, much smaller steps in dl are observed upon cooling, which occur also at much lower temperatures. When the humidity of equilibration at room temperature has been 86%, no drastic change in dl is seen upon temperature variation and dl shows values of about 54.0 Å. The same or very closely the same low-temperature limiting values of dl (near 53.5 Å) are reached below about 240 K by all samples equilibrated at room temperature at r.h. g 86%. The “dry” sample (equilibrated at 0% r.h.), on the other hand, has a lamellar lattice constant of only 49.6 Å, i.e., smaller by about 4 Å, with very little variation as a function of temperature. The pronounced decrease in lamellar spacing during cooling the hydrated purple membranes below Tfh is obviously due to the loss of water that had been intercalated between membranes at room temperature. The swelling seen upon reheating indicates that the water is returning into the inter-lamellar space. What happens to the departing water is immediately understood from the observation that Bragg peaks of crystalline ice are observed (data not shown here, see ref 71) as soon as the temperature falls below Tfh; they disappear when T is increased beyond Tmh. Freezing of water (D2O) is found to occur suddenly at temperatures between 268 and 266 K for the 100% and 98% samples and near 245 K for the 94% sample, i.e., well below the freezing point of bulk water (D2O) in these cases. The supercooling of inter-membrane water is more pronounced at lower initial hydration levels. Melting, on the other hand, occurs as a continuous transition in a temperature range starting at T ) Tmh*, i.e., already below the membrane water “freezing point Tfh”, and extending up to the membrane water “melting point Tmh”, which is above Tfh. No ice reflections are observed, when the r.h. of equilibration

is e86%. On the other hand, there is still water in the interbilayer space, since the lamellar spacing in this case is larger by about 4 Å than that of vacuum-dried samples. The practical absence of Bragg reflections suggests that this water is essentially noncrystalline. This water layer, with a minimum thickness of about 4 Å, consists of approximately two continuous monolayers of water, one belonging to each membrane surface. In addition to water of this layer, water in the slightly grooved lipid areas of the membrane presumed to be bordered by protruding protein loops of up to about 4 Å height,72 some water molecules penetrating into the membrane around lipid headgroups and the few water molecules bound in the interior of the protein34-36,46,47 represent the so-called “nonfreezing water” component much discussed in the literature.2,4 These results do not tell us unambiguously to what extent this water is still in the liquid or rather in a glassy state (“solid amorphous water” or “amorphous ice”) below 240 K. It is only possible to state that this component does not spontaneously transform into the crystalline state. The most likely reason for this is that the vapor pressure of this water component at T e 240 K is practically equal to that of ice at the same temperatures. A behavior qualitatively very similar to that discovered in purple membranes has also been observed in multilayer stacks of disk membranes obtained from bovine retinae.73 Disk membrane samples were equilibrated at 98% and 0% r.h., respectively. For comparison with corresponding results from purple membranes, the temperature dependence of dl for both disk membranes and purple membranes is shown in Figure 5. Quantitatively, larger values of dl for disk membranes as compared to purple membranes were measured. For dried samples we find dl-values of 59.2 Å for disk membranes and 49.1 Å for purple membranes, which corresponds to the

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J. Phys. Chem. B, Vol. 103, No. 38, 1999 8043 The above-described observations clearly demonstrated that the amount of water (corresponding to a certain hydration level) which remains closely attached to the biomolecule (often called “hydration water”) is a function of temperature. This fact is relevant for many investigations on dynamical and functional properties of biomolecules studied with hydrated powder samples or films of hydrated membrane sheets (see also Sections 4.2 and 5).

Figure 5. Lamellar spacings dl of (a) hydrated disk membranes (h ) 0.51) and (b) purple membranes (h ) 0.42) samples measured as a function of temperature (cooling: open downsided triangles; heating: solid upsided triangles). To obtain the water layer thickness (at room temperature: drt, below 240 K: dc) lamellar spacings of dried samples have been measured. Because these values do not show a temperature dependence (see Figure 4), they are represented by a straight dotted line.73

thickness of a sole single bilayer in each case.46,74 This difference is certainly due to the fact that rhodopsin is a larger molecule than bacteriorhodopsin, with much larger loop regions protruding into the inter-membrane space. The temperature dependence of the water layer thickness (dl (wet) - dl (dry)), characterized by the low-T limit dc, the room-temperature value drt, and the freezing transition temperature Tfh, is rather similar for purple membranes and disk membranes (see Figure 5). This comparison implies that both (rather different) types of membranes show a very similar interaction with the water molecules. The similarity of these phenomena observed in the case of purple and disk membranes and analogous results from pure lipid membranes75 suggest that we are dealing with a rather general property of lipid and protein-lipid membranes, namely the phenomenon of the temperature-dependent dehydration and rehydration of such membranes, which is caused by the presence of hydration forces and by the specific temperature dependence of the chemical potential of water.

4. Hydration-Induced Internal Molecular Motions 4.1. Dynamical Properties of Protein-Lipid Complexes. The dynamical features of membrane proteins and lipids in the biomembranes and the impact of hydration on the characteristics of the internal molecular motions were investigated by analyzing the scattering amplitudes of quasielastic and elastic components as measured with biomembrane samples at different hydration levels and different temperatures. The scattering amplitudes A0 and An as well as the line widths Hn (see eqs 6 and 7) generally exhibit a Q dependence which may be described by specific theoretical models used for the analysis of the measured data. Usually it is possible to derive the analytical incoherent scattering function only from relatively simple theoretical models, such as jump diffusion between different sites, diffusion in restricted volumes, or long-range translational diffusion (see, for example, refs 58,60). These models do not provide a detailed description of the dynamical behavior in very complex structures such as biological macromolecules. Too many molecular subunits (e.g., polypeptide side groups) perform their individual motions with different geometries and on very different time scales. They do, however, allow a phenomenological analysis of tof spectra measured with biological macromolecules. In this procedure, it is possible to fit the spectra with a certain number of Lorentzians and one elastic component, without using a specific model. The number of Lorentzians required within a given energy transfer range provides information about the complexity of the observed dynamics. The Q dependence of the incoherent structure factors (A0, An) and the line widths (Hn) give further information about properties of the motions (longrange translational diffusion, diffusion in restricted volumes, jump diffusion between discrete sites, etc.). As shown in the following subsections, this approach is very practical for studying dynamical properties in dependence of environmental conditions, such as temperature and hydration. For purple membrane and disk membrane samples, hydrated with D2O and measured at room temperature with different elastic energy resolutions, the phenomenological fit revealed the following general dynamical properties.73,76 (i) Within the given energy transfer range (|pω| e 3 meV) all spectra can be fitted with one elastic and two quasielastic components. The first quasielastic line width (H1) is strongly correlated with the resolution width (factor of 2 or 3 larger than Γres). The second line width (H2) is much broader, with values of a few meV (see Figure 6). Although solvent scattering represents only a few percent of the total scattering, it was separated from scattering of the membranes by applying a third quasielastic component in the fit. The line width (H3 ) 60 µeV) and the weight of this component were approximated on the basis of studies on the solvent.23,67 The weight A3 is a function of the ratio of scattering power between D and H (SD/SH ) 1/11) and the hydration level of the sample h. It can be approximated by A3 ) h × 1/11; e. g., for h ) 0.4 g of D2O /g of purple membranes the resulting A3 ) 0.036 (Figure 6). (ii) The quasielastic incoherent structure factors (A1, A2) increase from small to large Q values. The line widths obtained do not show significant Q dependences, indicating localized motions (line widths as a function of Q are


Figure 6. The fit of a spectrum as measured at room temperature with a hydrated (h ) 0.4 with D2O) purple membrane sample is shown to demonstrate what amount the elastic and the different quasielastic components contribute to the total scattering. The spectrum was measured at the multichopper time-of-flight spectrometer NEAT located at the Hahn-Meitner Institut, Berlin78 using a wavelength of λ ) 5.1 Å which corresponds to an elastic energy resolution of Γres ) 100 µeV (fwhm). This spectrum was obtained by grouping all spectra measured in a range of scattering angles φ from 13.3° to 136.7° with a weighted mean of φmean ) 55.4°. The experimental points were fitted by the total scattering function composed of one elastic (resolution) and three quasielastic Lorentzian-shaped contributions (L1-L3). The figure legend in (b) is also valid for (a). The quasielastic scattering of L1 and L2 correspond to internal motions of bacteriorhodopsin and the lipids. The much smaller component of L3 represents local motions of the D2O solvent. The shape of the elastic scattering was determined by vanadium measurements. The lower part of this figure (b) shows an enlargement of the quasielastic scattering. Top of the frame in (b) corresponds to 10% of the elastic peak intensity in (a). Note the different energy transfer ranges in (a) and (b).

not shown here, see refs 24,73,77). (iii) Because slower motions are resolved only by measurements with higher energy resolution (e.g., Γres ) 100 µeV (τ e 6 ps) and Γres ) 34 µeV (τ e 20 ps) [fwhm]) the integral of the total quasielastic incoherent scattering increases with decreasing Γres. These results indicate that the major part of the scattering is due to local diffusive motions of hydrogen atoms within restricted volumes (with a diameter of a few angstroms) and with correlation times from 0.1 up to 100 ps. The fact that we

Fitter et al.

Figure 7. A comparison of spectra measured with hydrated purple membranes and disk membranes. Both samples were measured at room temperature (T ) 293 K) with the NEAT spectrometer using λ ) 6.2 Å (Γres ) 34 µeV [fwhm]). Although all spectra were fitted with three quasielastic components within an energy transfer range of |pω| e 3 meV, the components and the resulting values (An, Hn) of only two components (L1 and L2, see legend Figure 6) are shown here. The spectra presented were grouped to φmean ) 55.4°. The largest intensity values of the spectra (top of the frame), correspond to 10% of the elastic peak intensity in each case.

obtain different line widths, even for only slightly different energy resolution, is a strong indication that the quasielastic spectra of purple membranes and disk membranes are characterized by a much larger number (larger than two) of Lorentzians showing a broad and more or less continuous distribution of line widths. The line widths obtained in the phenomenological fit at specific energy resolutions must be understood as mean values of a distribution of line widths, and the related motions are characterized by the corresponding range of different correlation times.73,76 A comparison of the dynamical behavior as obtained from hydrated purple membranes and disk membranes helps to reveal some general features of internal motions occurring in a protein-lipid complex (see Figure 7). Both membranes have been hydrated at 98% r.h. D2O (which led to corresponding h ) 0.42 for purple membranes and to h ) 0.51 for disk membranes) and the interaction of solvent with the membranes is rather similar for both types of membranes as shown in Figure

Feature Article 5. [A direct comparison of h-values between purple membrane and disk membrane samples is not straightforward because both membranes have different membrane thicknesses, different protein-lipid ratios, and therefore different specific densities (see Table 1 and Figure 5).] The quasielastic structure factors (A1 and A2 given in Figure 7) are 10-20% larger for the disk membrane sample as compared to the purple membrane sample with respect to the given energy resolution of Γres ) 34 µeV. Results of spectra measured with Γres ) 100 µeV (data not shown here) revealed a similar difference between both membranes. The reason for the increased internal flexibility of disk membranes as compared to purple membranes seems to be related to different membrane protein structures of rhodopsin and bacteriorhodopsin and to the rather different composition of both membranes. Larger, and maybe more flexible, loop regions of rhodopsin as compared to bacteriorhodopsin35,50 may contribute to larger quasielastic structure factors as found for disk membranes. But a much stronger influence on the dynamics seems to have the much larger content of lipids in disk membranes as compared to purple membranes (see Table 1), which is obviously responsible for a larger contribution of internal motions on the picosecond time scale. The particular role of lipids for dynamical properties in biological membranes will be discussed in more detail in Section 4.3. 4.2. Dynamical (Glasslike) Transition. The temperature dependence of incoherent neutron scattering from purple membrane samples, hydrated at h ) 0.4 g of D2O per g of purple membrane, was studied using a moderate energy resolution of Γres )100 µeV (fwhm). Phenomenological fits (see, for example, Figure 6) of spectra, measured with samples which were cooled from 297 to 10 K, yielded the following results. (i) It was possible to fit all spectra with two quasielastic components (H1 ) 120 µeV, τ1 ) 5.5 ps; H2 ) 2 meV, τ2 ) 0.5 ps). These components, with linewidths kept fixed over the whole temperature range, are characterized by temperaturedependent structure factors (for more details see ref 76). (ii) The temperature dependence of the incoherent structure factors A1 (Figure 8a) indicates, that the narrow component (H1 ) 120 µeV) includes only stochastic motions. Below a temperature Td ∼ 200 K this component does not exist and we observe a dynamical transition characterized by the onset of the these slower “large-amplitude” motions above Td. (iii) The faster motions (represented by A2 with H2 ) 2.0 meV) show a significant amplitude even at low temperatures increasing more or less linearily with T up to approximately 200 K (Figure 8b). This linear T dependence is typical for vibrational motions often described by Debye-Waller factors. Above T ≈ 200 K, we observe a deviation from this linear T dependence with an additional increase at higher temperatures. Such a behavior indicates that this component includes only vibrational motions at low temperatures and additional stochastic “large-amplitude” motions above Td. (iv) A further “transition” is observed for the slower motions at temperatures between 250 and 270 K (Figure 8a). As known from neutron diffraction studies71 (see Section 3.2) this additional decrease of internal flexibility is due to membrane dehydration upon solvent freezing. The main feature of the investigated temperature dependence is a drastic increase of internal molecular motions above Td as compared to low temperatures. This phenomenon is the wellknown dynamical “glass transition”, which has been observed for various biomolecules previously in many studies mainly on water-soluble globular proteins61,79-81 but also on purple membranes82 with Td ) 170-230 K. According to a model


Figure 8. The temperature dependence of quasielastic incoherent structure factors for “slow” motions (a) and “fast” motions (b) as obtained from phenomenological fits of spectra measured with hydrated purple membrane samples (h ) 0.4). The solid line in (b) represents a linear decrease in A2 at temperatures below 200 K, indicating vibrational motions in this temperature regime (for more details, see ref 76).

introduced by Frauenfelder and co-workers,61 the stochastic “large-amplitude” motions are related to “barrier crossing” between conformational substates. In this picture, the ability to perform transitions between different conformational substates, which is supposed to be an important property of a functional biomolecule, is possible only at temperatures above Td. Many investigations on proteins with different kinds of solvents (e.g., solvent mixtures water/trehalose) and different amounts of solvent indicate a strong influence of solvent and solvent properties on the characteristics of the dynamical transition.83-86 Therefore additional temperature-dependent measurements on purple membrane samples with different hydration levels have been performed in order to study the influence of solvent on the dynamical behavior of the protein-lipid complex. The temperature dependence of slow stochastic “large-amplitude” motions of a moderately hydrated (h ) 0.38), a weakly hydrated (h ) 0.18), and a dried (h ) 0.06) sample is shown in Figure 9a. Within the limits of error, the dynamical transition occurs at the same temperature. A1 is rather similar in the lowtemperature regime up to T ) 250 K for all hydration levels. Above a temperature of T ) 250 K the quasielastic incoherent structure factor A1 is larger for higher hydration levels. Interest-


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Figure 10. Quasielastic incoherent structure factors A1 (“slow” motions; solid triangles) and A2 (“fast” motions; open circles) determined from purple membrane samples measured at room temperature as a function of hydration level.76

Figure 9. (a) Temperature dependence of the quasielastic incoherent structure factor A1 of purple membrane samples with different hydration levels: 98% r.h. (h ) 0.38), 86% r.h. (h ) 0.18), 0% r.h. (h ) 0.06). (b) A2 values, representing faster motions, are given for sample hydrations h ) 0.38 (solid triangles) and h ) 0.18 (open squares).

ingly, even the dried sample shows a significant amplitude of A1 [7% ((2%) of the total scattering] at room temperature and a transition near 200 K. In contrast to this, dried powders of globular proteins (dried R-amylase with h ) 0.05) revealed a much smaller contribution of A1 (2% of the total scattering) which does not vary within the limits of error ((2%) over the temperature and shows therefore no dynamical transition.87 The amplitude of A2, which is related to faster motions, shows significantly larger values for weakly hydrated (h ) 0.18) samples than that of moderately hydrated (h ) 0.38) samples below a temperature of about 270 K (Figure 9b). This reduced vibrational component in more-hydrated samples as compared to less-hydrated samples at low temperatures is certainly related to solvent properties; either the partially crystallized solvent inhibits the vibrational motions, or the solvent is still able (even at low temperatures) to damp those vibrational motions. In the latter case, the damped motions appear too slow (with τ > 6 ps which is not resolved by the given energy resolution of Γres ) 100 µeV) to occur in the quasielastic contribution of A1. The generality of this phenomen can be appreciated from a comparison of purple membrane data with data obtained from wet and dry samples of hemoglobin88 and of R-amylase,87 where a

similar behavior of low-temperature vibrational motions was observed. On the basis of additional measurements performed at room temperature and using samples hydrated at various hydration levels (Figure 10), a comparison of A1 and A2 revealed that slower motions are more strongly influenced by the hydration level as compared to faster motions. In addition to studies on hydrated purple membranes, measurements on hydrated disk membranes were also performed as a function of temperature. Qualitatively, the samples of disk membranes show the same behavior as compared to purple membranes with a dynamical transition at Td ∼ 200 K. These measurements were performed with disk membranes, which were first cooled and subsequently heated. Interestingly, the temperature dependence of A1 shows a pronounced hysteresis similar to dl values given in Figure 5 (see also ref 87). 4.3. The Role of Lipids in the Hydration Process. To study the influence of lipids on the dynamical behavior of bacteriorhodopsin we performed measurements with natural purple membranes composed of 75% bacteriorhodopsin (w/w) and 25% lipid (w/w) and with partially delipidated purple membranes having only 5% lipid (w/w).89 Both types of membranes have been hydrated at 75% r.h. (h ) 0.15) and at 98% r.h. (h ) 0.4) (in D2O atmosphere). As an alternative procedure to the phenomenological fit, the data have been analyzed with a simple model, which should reproduce some general properties of the complex dynamical behavior. It describes continuous diffusion inside the volume of a sphere with the radius r, where the EISF is given by90

[

A0(Q) ) 3

]

sin(Qr) - (Qr)‚cos(Qr) (Qr)3

2

(14)

Because our data are limited in momentum transfer Q (with |Qmax| ) 2.25 Å-1) and we used only an energy transfer range with |pω| e 0.8 meV, we need to consider only the first quasielastic component (see eq 6). Therefore, the theoretical scattering function is only parametrized by r (determining A0 and A1 ) 1 - A0) and the width of the quasielastic contribution H1 ) (τ1)-1. These parameters must be understood as values averaged over all the motions observable in the time window

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J. Phys. Chem. B, Vol. 103, No. 38, 1999 8047 TABLE 2: Results from the Model Fit (local diffusion inside a sphere with radius r, see eq 14) for Native and Delipidated Purple Membrane (PMnat, PMdelip) Samples at Two Different Hydration Levelsa

radius [Å] 〈u2〉 [Å2]

PMdelip (98%r.h.)

PMnat (98% r.h.)

PMdelip (75% r.h.)

PMnat (75% r.h.)

0.57 0.052

0.77 0.096

0.45 0.076

0.52 0.130

a

The parameter r represents an “amplitude” of slow stochastic motions and the averaged mean square displacements 〈u2〉 (as obtained from Debye-Waller factors, see eq 6) are related to amplitudes of fast vibrational motions.73

Figure 11. Q-dependence of elastic incoherent structure factors as obtained at two different hydration levels for native and delipidated purple membranes. The experimental EISF (A0) (solid symbols) as obtained from fits are shown as a function of momentum transfer Q. For this purpose, all 16 spectra have been fitted separately with radius r as a free parameter. The statistical error of the experimental EISF is approximately (0.015, which is about the size of the symbols. The corresponding theoretical EISF (PMnat: dashed-dotted line, PMdelip: solid line) were obtained from model fits (according to eq 14 with resulting r values given in Table 2), where all spectra have been fitted simultaneously (for details see ref 89).

of our experiments (a few picoseconds). The radii r, determining the volume of stochastic localized motions, can be interpreted as “amplitudes” of these motions. These “large-amplitude” motions give the main contribution to what can be called “internal flexibility”. The averaged “mean square displacements” 〈u2〉 represent amplitudes of mainly vibrational motions. All spectra show a Q-independent line width of H1 ) 150 µeV, which corresponds to a correlation time of τ1 ) 4.4 ps. The comparison of internal molecular motions occurring in natural and in delipidated purple membranes revealed the following results (see Figure 11 and Table 2):

We find more internal flexibility, related to more quasielastic scattering with larger radii r and to larger 〈u2〉, in natural purple membranes as compared to delipidated purple membranes (see Table 2). As shown in Figure 11, a steeper decrease of the A0 with Q is related to larger “amplitudes” r and therefore to a larger internal flexibility. The difference in flexibility between natural and delipidated purple membranes is relatively weak in the case of weakly hydrated samples (75% r.h., h ) 0.15) and more pronounced in the case of strongly hydrated samples (98% r.h., h ) 0.4). With respect to the obtained radii r, and as already known from previous studies, 98%-samples exhibit more internal flexibility than 75%-samples. This effect is more pronounced in the case of natural as compared to delipidated purple membranes. Due to damping of the vibrational motions in 98%samples, the mean square displacements 〈u2〉 are smaller as compared to 75%-samples. This kind of h-dependent “solvent damping” was also found in the phenomenological analysis of purple membranes (see A2 in Figure 9b). These results indicate that either the lipids themselves are more flexible than the membrane protein bacteriorhodopsin (a high lipid content leads to an increased “overall” internal flexibility in the purple membrane as compared to a low lipid content), or that more lipids increase the flexibility of the membrane proteins. A further, and most probable, possibility is a combination of both influences. It is reasonable to assume that in the tightly packed protein-lipid complex the dynamics of bacteriorhodopsin is coupled with the dynamics of the lipids. The fact that the differences in the flexibility between natural and delipidated purple membranes are much more pronounced in 98%-samples, gives strong evidence that mainly the presence of hydration water, which is attached to the polar lipid headgroups at high hydration levels, increases the internal flexibility of the protein-lipid complex.89,91 This hypothesis is supported by diffraction experiments performed with natural purple membranes, where at high hydration levels (98% r.h.) a larger in-plane cell parameter ds was found (62.4 Å) as compared to 75%-samples (61.4 Å). The reduction of the in-plane cell dimension is related to a removal of solvent molecules which were located around the lipid headgroups.46,66 Although, on the basis of the present data, we are not able to distinguish between dynamics of the lipids and dynamics of bacteriorhodopsin, we can draw the conclusion that the amount of lipids and their ability to attract solvent molecules is strongly related to “largeamplitude” motions of the whole purple membrane. This explanation seems to be also valid under consideration of the observed difference in the dynamical behavior between the natural purple membrane and the disk membrane as presented in Section 4.1. All above-presented studies (Sections 4.1, 4.2, 4.3) revealed significant effects of temperature, hydration, and protein/lipid ratios on the dynamical behavior. One of the main results which


Figure 12. Time constant characterizing the absorption signal of the decay of the M-intermediate in natural purple membranes (squares) and in delipidated purple membranes (circles) as a function of relative humidity and measured at room temperature (for more details, see ref 89).

comes out of these studies seems to be the following: A reduced internal flexibility of biomembranes, due to lowering the temperature or because of reducing the lipid content in membranes, is caused to a large extent by dehydration which is accompanying these processes. More or less mobile water molecules intercalated between lipid headgroups and attached to the membrane surface seem to provide the “driving force” for thermal stochastic “large-amplitude” motions. The particular relevance of these motions for the (biological) function of the purple membrane will be discussed in the next section. 5. Influence of Hydration on the Function of Bacteriorhodopsin The impact of the hydration process on the functional aspects of biomembranes has been investigated in many studies for the proton pump bacteriorhodopsin in the purple membrane (see, for example, refs 39,47). With respect to the function of bacteriorhodopsin in the purple membrane, it is necessary to distinguish between at least three different categories of water: (i) 2-3 water molecules which were found in the vicinity of the Schiff base at the protonated linkage of the chromophore and the amino acid lysine-216 of the protein moiety, (ii) a string of water molecules, located in the interior of the protein structure, which is supposed to act as a kind of “proton wire” and accomplish the proton transfer across the bacteriorhodopsin molecules,46,47,92 and (iii) hydration water which is located inside the protein, at the protein surface (e.g., loop regions), intercalated between lipid headgroups or attached to the membrane surface. This hydration water is supposed to be essential for the proton migration along the membrane surface and required for an internal flexibility of the whole membrane on a fast picosecond time scale to “lubricate” light-induced conformational changes on a millisecond time scale.39 The functional importance of water molecules for bacteriorhodopsin is clearly demonstrated in Figure 12 (see solid symbols for natural purple membranes), illustrating the dependence of the decay rate of the intermediate M (functionally important

Fitter et al. intermediate state of the bacteriorhodopsin photo- and pumpingcycle, see, for example, ref 1) on the relative humidity. Both, the decay of M and the regeneration of the ground state are strongly slowed below 80% r.h. As measured with optical pHindicators, the proton pumping activity is decreased below 70% r.h. and no activity is observed below 50% r.h., although there is still a very slow photocycle.47 The photocycle and the proton pumping cycle of bacteriorhodopsin are accompanied and most probably propelled by structural changes in both the chromophore retinal and the protein. The light-induced appearance of tertiary structural changes in the protein, predominantly at helix F and G, has been established by various investigations using neutron38 and time-resolved X-ray diffraction.39,93 At the same threshold value of about 60-50% r.h., where proton transport stops, also the light-induced conformational changes in the protein cease.39 All these properties of bacteriorhodopsin, i.e., the photocycle, the proton pumping cycle, and the conformational changes, are influenced by the state of hydration, proving the functional importance of water. Here we want to discuss functional properties of BR which are related to aspects of the hydration process presented in Sections 3 and 4. In the context of energy conversion processes, it is not only of interest how protons are pumped across the membrane (to be accomplished by bacteriorhodopsin) but also how protons diffuse along the membrane surface to reach other proteins (for example, the H+-ATP synthase as a proton consumer). Recent studies, using optical proton sensors (fluorescein and pyranine), revealed that proton migration along the membrane surface is much faster than the release of protons from the membrane surface into excess water.44 This indicates that long-range proton migration along the membrane surface might be accomplished mainly by the first water layer which is in direct contact with the membrane surface. As shown in Section 3.2 the binding of the first water layer to the membrane surface is rather strong and seems to represent a kind of “border” to the excess water with respect to fast proton conduction. Whether the self-diffusion of inter-membrane water (or part of this water) is relevant for the proton migration along the membrane surface is still an open question. Instead of a water-assisted proton-transfer process, on the basis of diffusing water molecules acting as vehicles alternating between an acceptor state (forming H3O+ ions) and a donor state (to give OH- ions), more immobilized water molecules of the first hydration layer might play the central role in the proton diffusion. Possibly this, probably dynamically restricted (mainly rotational but little translational motions of the water) layer of hydration water, which represents a twodimensional H-bonded network, allows fast proton diffusion. With higher rotational mobility of water molecules parallel to the membrane, the reduced dimensionality (only two dimensions instead of three with respect to a certain time scale) of the H-bonded network seems to allow an increased lateral diffusion of protons. As already mentioned above, the specific function of bacteriorhodopsin (or of a biomolecule in general) is not only determined by structural features but also by dynamical properties. In particular, a certain amount of “stochastic largeamplitude” motions is required for a proper function, here for the light-induced proton pumping of bacteriorhodopsin. As a demonstrating example for this hypothesis, both the dynamical behavior and functional parameters were studied. In the case of natural and delipidated purple membrane, time constants characterizing the bacteriorhodopsin photocycle have been compared to results of the dynamical properties as a function of hydration level (see Section 4.3). Time constants of the

Feature Article spectroscopic M-intermediate, which are related to the M-decay and to proton uptake, are significantly larger in “dried” and/or delipidated purple membranes as compared to “wet” and natural membranes (see Figure 12). As a demonstrative confirmation of the hypothesis, less-efficient proton pumping (indicated by a prolonged M-decay) was found to be accompanied by a reduced internal flexibility of the purple membrane.89 A similar correlation was also observed for temperature-dependent measurements of the M-decay and of the internal dynamics. At low temperatures, where the internal flexibility is reduced,76 the M-decay is significantly prolonged.94,95 6. Concluding Remarks Many studies on the purple membrane as an important prototype (or model system) of a biological membrane and the results of neutron scattering studies presented here demonstrated how hydration water and various aspects of the hydration process play a central role for the essential properties of the membrane and for the function of bacteriorhodopsin. Beside particular aspects of the purple membrane, e.g., the location of water molecules in the interior of the bacteriorhodopsin structure which is supposed to be relevant for the transversal proton translocation, the interaction of water molecules with the membrane surface and with specific sites inside the membrane seems to be a more general phenomenon in the hydration process of biological membranes. With respect to the internal flexibility of molecules of the membranes on a picosecond time scale, h-dependent measurements demonstrated that these motions are “lubricated” by water molecules. Although various similarities in the hydration process were found for biomembranes and for water-soluble proteins, the presence of lipids in biomembranes seems to be the reason for some noticeable differences in the hydration process. The comparison of membrane stacks and powders of globular proteins revealed a small but pronounced component of “large-amplitude” stochastic motions in dried membrane stacks, while these motions are more or less absent in dried powders of water-soluble proteins.87 The data indicate that it is not possible to “dry out” the “large-amplitude” stochastic motions completely in biomembranes, which is on the other hand possible in the case of protein powders. The particular geometry of membranes and the strong interaction of lipids with water molecules seem to be the reason for this unique property as compared to other biological macromolecules, such as water-soluble proteins or nucleic acids. With respect to a particular molecular mass of a biomembrane fragment and of a globular protein, the same amount of water in both types of samples is required to hydrate a smaller surface area of planar membranes as compared to a larger surface area of more or less spherical proteins. Beside often functionally important (“material”) properties of the hydration water and of biomembranes as revealed by the presented neutron scattering studies, it is important for future experiments to extend our knowledge about how these properties are related to the function of the membrane or the membrane proteins. Bacteriorhodopsin in purple membranes and rhodopsin in disk membranes offer the possibility to perform measurements with the membrane proteins in different structural conformations. Up to now the proton pump bacteriorhodopsin and the photoreceptor rhodopsin have been studied by incoherent neutron scattering mainly in the ground state. To elucidate the role of hydration properties for the function, more ambitious measurements with light-activated intermediate states are required. We have already performed measurements in this direction with both membrane proteins.96 To decide whether INS is really applicable

J. Phys. Chem. B, Vol. 103, No. 38, 1999 8049 to detect dynamical features which are related directly to the function, future investigation need to be supported by further methodical developments. Acknowledgment. The technical staff of the neutron facilities at the Hahn Meitner Institut (Berlin, Germany), at the Institute Laue-Langevin (Grenoble, France), and at the ISISfacility (Didcod, GB) are acknowledged for their assistance during the measurements. We are obliged to K. P. Hoffman and O. P. Ernst for their cooperation in the project on disk membranes, to Th. Hauss for cooperation on diffraction studies, to S. A. W. Verclas for time-resolved absorption spectroscopy measurements, and to G. Bu¨ldt for valuable discussions. This work was supported by Grant 03-DE5DA1-8 from Bundesministerium fu¨r Bildung und Forschung (to J.F. and N.A.D.), by the Deutsche Forschungsgemeinschaft (SFB 472 to N.A.D.), and by Fonds der chemischen Industrie (to N.A.D.). References and Notes (1) Stryer, L. Biochemistry, 4th ed.; W. H. Freemann and Company: New York, 1995. (2) Kunz, I. D.; Kauzman, W. AdV. Protein Chem. 1974, 28, 239. (3) Finney, J. L.; Poole, P. L. Comments Mol. Cell. Biophys. 1984, 2, 129. (4) Rupley, J. A.; Careri, G. AdV. Protein Chem. 1991, 41, 37. (5) Byrant, R. G.; Shirley, W. M. Biophys. J. 1980, 32, 3. (6) Otting, G.; Liepinsh, E.; Wu¨thrich, K. Science 1991, 254, 974. (7) Densiov, V. P.; Halle, B.; Faraday Discuss. 1996, 103, 227. (8) Steinhoff, H. J.; Kramm, B.; Hess, G.; Owerdieck, C.; Redhardt, A. Biophys. J. 1993, 65, 1486. (9) Kossiakokoff, A. A.; Sintchak, M. D.; Shungin, J.; Presta, L. G. Proteins 1992, 12, 223. (10) Schoenborn, B. P.; Garcia, A.; Knott, R. Prog. Biophys. Mol. Biol. 1995, 64, 105. (11) Teeter, M. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 6041. (12) Levitt, M. Ann. N.Y. Acad. Sci. 1981, 367, 162. (13) Brooks, C. L.; Karplus, M. J. Mol. Biol. 1989, 208, 159. (14) Steinbach, J. P.; Loncharich, R. J.; Brooks, B. R. Chem. Phys. 1991, 158, 383. (15) Knapp, E. W.; Muegge, I. J. Phys. Chem. 1993, 97, 11339. (16) Bizarri, A. R.; Wang, C. X.; Chen, W. Z.; Cannistraro, S. Chem. Phys. 1995, 201, 463. (17) Miyazaki, Y.; Matsuo, T.; Suga, H. Chem. Phys. Lett. 1993, 213, 303. (18) Green, J. L.; Fan, J.; Angell, C. A. J. Phys. Chem. 1994, 98, 13780. (19) Doster, W.; Bachleitner, A.; Dunau, R.; Hiebl, M.; Lu¨scher, E. Biophys. J. 1986, 50, 213. (20) Middendorf, H. D. Annu. ReV. Biophys. Bioeng. 1984, 13, 425. (21) Bellissent-Funel, M.-C.; Teixera, J.; Bradley, K. F.; Chen.; Crespi, H. L. Physica B 1992, 180/181, 740. (22) Ko¨nig, S.; Bayerl, T. M.; Coddens, G.; Richter, D.; Sackmann, E. Biophys. J. 1995, 68, 1871. (23) Lechner, R. E.; Dencher, N. A.; Fitter, J.; Dippel, Th. Solid State Ionics 1994, 70/71, 296. (24) Fitter, J.; Lechner, R. E.; Bu¨ldt, G.; Dencher, N. A. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 7600. (25) Sartor, A.; Hallbrucker, A.; Mayer, E. Biophys. J. 1995, 69, 2679. (26) Reid, C.; Rand, R. P. Biophys. J. 1997, 72, 1022. (27) Israelachvili, J.; Wennerstroem, H. Nature 1996, 379, 219. (28) Finney, J. L. Faraday Discuss. 1996, 103, 1. (29) Acrangeli, C.; Bizzarri, A. R.; Cannistraro, S. Chem. Phys. Lett. 1998, 291, 7. (30) Nengendra, H. G.; Sukumar, N.; Vijayan, M. Proteins 1998, 32, 229. (31) Mitchell, P. Nature 1961, 191, 144. (32) Henderson, R.; Baldwin, J. M.; Ceska, T. A.; Zemlin, F.; Beckmann, E.; Downing, K. H J. Mol. Biol. 1990, 213, 899. (33) Kimura, Y.; Vassylyev, D. G.; Miyazawa, A.; Kidera, A.; Matsushima, M.; Mitsuoka, K.; Murata, K.; Hirai, T.; Fujiyoshi, Y. Nature 1997, 389, 206. (34) Pebay-Peyroula, E.; Rummel, G.; Rosenbusch, J. P.; Landau, E. M. Science 1997, 277, 1676. (35) Essen, L.-O.; Siegert, R.; Lehmann, W. D.; Oesterhelt, D. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 11673. (36) Luecke, H.; Richter, H.-T.; Lanyi, J. K. Science 1998, 280, 1934. (37) Stoeckenius, W.; Lozier, R. H.; Bogomolni, R. A. Biochim. Biophys. Acta 1979, 505, 215.

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