5.6 Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

5.6 Atomic Force Microscopy and Electron Microscopy of Membrane Proteins A Engel, Case Western Reserve University, Cleveland, OH, USA; Maurice E. Mu¨ller University of Basel, Switzerland r 2012 Elsevier B.V. All rights reserved.

5.6.1 Introduction 5.6.2 Electron-crystallography 5.6.2.1 2-D-crystallization 5.6.2.2 Sample Preparation 5.6.2.3 Image Formation 5.6.2.4 Electron Diffraction 5.6.2.5 Data Acquisition 5.6.2.6 Data Processing 5.6.2.7 Examples of Structures Solved by E-crystallography 5.6.2.7.1 Bacteriorhodopsin 5.6.2.7.2 Aquaporins 5.6.3 Atomic Force Microscopy 5.6.3.1 Instrumentation 5.6.3.2 Sample Preparation 5.6.3.3 Data Acquisition 5.6.3.4 Image Processing 5.6.3.5 Insights from AFM Topographs 5.6.3.5.1 Bacteriorhodopsin 5.6.3.5.2 E. coli Outer Membrane Porin OmpF 5.6.3.5.3 Native Membranes 5.6.4 Single Molecule Force Spectroscopy 5.6.4.1 Instrumentation and Methodology 5.6.4.2 Sample Preparation 5.6.4.3 Data Acquisition 5.6.4.4 Data Processing 5.6.4.5 Insights from Single Molecule Force Spectroscopy (Examples) 5.6.4.5.1 Visualizing the Site where a Membrane Protein was Unfolded 5.6.4.5.2 Localizing Unfolding Barriers 5.6.4.5.3 Localizing a Ligand Binding Site 5.6.4.5.4 Unfolding from Different Ends 5.6.4.5.5 Refolding Back into the Bilayer 5.6.5 Perspectives Acknowledgments References

Abbreviations 2-D 2-DFT 3-D 3-DFT AQP bR CCD CMC CTF DLVO EM FM

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two-dimensional two-dimensional Fourier transform three-dimensional three-dimensional Fourier transform aquaporin bacteriorhodopsin charge-coupled device critical micellar concentration contrast transfer function Derjaguin, Landau, Verwey, Overbeek electron microscopy frequency modulation

HOPG LPR MBCD PDB PEG PSD psf SD SECM SMFS SNR WLC

93 94 94 94 95 97 98 98 100 100 100 103 103 105 105 106 107 107 107 109 110 110 111 111 113 113 113 113 113 114 115 115 116 116

highly oriented pyrolytic graphite lipid-to-protein ratio methyl-b-cyclodextrin Protein Data Bank polyethyleneglycol position-sensitive photodiode point spread function standard deviation scanning electrochemical microscopy single molecule force spectroscopy signal-to-noise ratio worm-like-chain

Comprehensive Biophysics, Volume 5

doi:10.1016/B978-0-12-374920-8.00511-7

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

Glossary Atomic force microscopy A method to measure the surface topography of proteins in solution. The deflection of a cantilever with a sharp tip at its end is recorded while the sample surface is raster-scanned below the tip. Bio-beads Polystyrene beads with small hydrophobic pores that allow detergent molecules to be adsorbed. Critical micellar concentration Concentration at which detergent molecules assemble spontaneously into micelles to bury the hydrophobic moiety in a hydrophobic core. Cyclodextrin Cyclic polysaccharide with a hydrophobic inner surface Detergent Amphiphilic molecule with limited water solubility as monomer, but highly soluble in the micellar form. Dissolution point The point in the solubilization process where a sufficient number of detergent molecules is available to solubilize all components of a biological membrane.

5.6.1

Introduction

Biological membranes enclose and compartmentalize cells of all organisms, acting as effective insulators and selective filters. They are composed of a phospholipid bilayer and integrated membrane proteins, which relay information or chemical substrates across the membrane and are key players in the biochemical events that take place either at the surface of cells or within membrane-bound organelles. Depending on the biological context, membrane proteins act as receptors, enzymes, channels, transporters, structural proteins or cell-cell adhesion molecules and, as such, contribute to an astounding variety of essential cellular functions, including transmembrane signaling, homeostasis, and energy conversion. To optimize their function, membrane proteins are clustered in domains that are restricted by the physical and chemical constraints of the lipid bilayer.1 Because of their central role in all physiological processes membrane proteins are important drug targets. While around 40% of all genes are known to encode membrane proteins,2 our understanding for this important class of proteins is growing only slowly, because of their highly hydrophobic nature and their intricate subunit structure. Prevalent structure determination tools, such as X-ray crystallography and solution NMR spectroscopy, which are increasingly successful for the structure determination of detergent-solubilized species, are at a distinct disadvantage when membrane proteins are embedded in a lipid bilayer. Because the detergent cannot duplicate the physical/chemical environment of the lipid bilayer, detergent solubilized membrane proteins are often unstable and tend to aggregate. Moreover, conditions that promote interactions between extramembraneous protein domains required for 3-D crystallization and at the same time preserve the protein’s hydrophobic core region are difficult to find. These problems are clearly reflected in the Protein Data Bank (PDB): out of B60 000 structures deposited as of July 2010, only B553 structures come from 204 different membrane proteins (http://blanco.biomol.uci.edu/

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Electron crystallography The primary method to determine the structure of 2-D crystals. Images as well as diffraction patterns are recorded at low electron-doses and keeping the sample at low temperature. Combining the information from many images and diffraction patterns taken at different projection angles allows the 3-D structure of a crystal to be determined at atomic resolution. Saturation point The point in the solubilization process where the biological membrane is saturated with detergent molecules intercalating between lipids and proteins. Solubilization A biological membrane is solubilized by interaction with a detergent. Ultimately individual components of the membrane, that is, the proteins and lipids, are incorporated in individual detergent micelles. Tag Engineered appendix of a recombinant protein with a specific structure suitable for affinity chromatography. Two-dimensional crystal Regular assembly of threedimensional molecules in a planar array.

Membrane_Proteins_xtal.html). Methods that allow membrane proteins to be assessed in the native membrane or when reconstituted in a lipid bilayer are thus of great interest. Electron crystallography is one attractive approach. This involves the imaging and diffraction of 2-D crystals in a transmission electron microscope as demonstrated by pioneering work on bacteriorhodopsin.3 Based primarily on early developments by Henderson and colleagues, electron crystallography has continued to be a powerful tool for studying the 3-D structure of membrane proteins at medium and highresolution, yielding several atomic structures (see ref. 4). To achieve this, highly ordered 2-D crystals are an indispensable prerequisite.5 Less well ordered crystals have allowed the 3-D structure of approximately 25 other unique membrane proteins to be determined at medium resolution (5–8 A˚), and continuing efforts are expected to produce atomic models in the near future. The recent structure of aquaporin-0 (AQP0) is not only of particular interest for its remarkably high resolution (1.9 A˚), but also for its unique ability to reveal the conformation of all the lipid molecules that surround the protein.6 Thus, despite advances in the application of solid state NMR and molecular dynamics simulations, electron crystallography represents the best approach for understanding membrane protein structure in the context of the lipid bilayer. Atomic force microscopy (AFM),7 the other powerful approach, offers unique capabilities for observing and manipulating single proteins in a physiological environment, and allows their function and dynamics to be assessed directly. The surface structure of membrane protein arrays can be determined by AFM in buffer solutions at a lateral resolution of 4–10 A˚ and a vertical resolution of 1–2 A˚.8 Such information can be obtained from regularly or randomly packed membrane proteins. The resolution of the method is determined by the size of the probe that touches the surface – ultimately it is the atom at the tip apex. Since the tip cannot contour deep and narrow crevasses properly, objects which are rather flat – like biological membranes with proteins that

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Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

protrude only by a some 10 A˚ – are quite amenable to highresolution imaging by AFM. Native and reconstituted membranes densely packed with proteins embedded in the lipid bilayer have given the most significant results, depicting the surface structure of single membrane proteins with a lateral resolution of 5 A˚ and a vertical one that is even better.9–11 Membrane proteins can also be addressed directly with the tip and unfolded, revealing the unfolding process of single proteins.8,12–14 The forces arising in such events may thus be studied with unparalleled resolution and sensitivity.12 The folding energy landscape can be analyzed both in magnitude but also terms of the dynamics of the response, and energy barriers can be assigned to specific residues. The binding of individual ligands may also be visualized, and the alterations in the mechanics of the proteins may be detected and localized again, at the precision of a few amino acids.12,15 This chapter provides information on the technical aspects and the limitations of these two powerful approaches, used for investigating the structure and function of membrane proteins embedded in a lipid double layer, and it presents typical applications.

5.6.2 5.6.2.1

Electron-crystallography 2-D-crystallization

Two-dimensional crystals consisting of membrane proteins and lipids can be produced in different ways. The first method involves the induction of regular packing of a highly abundant protein in its native membrane. This is achieved by eliminating interspersed lipids using lipases16 or by extracting lipids with specific detergents.17 Although this is the most gentle 2-D crystallization method, because it does not require solubilization of the membrane protein, it is not generally applicable. The second method reconstitutes the purified membrane protein into a lipid bilayer at high protein density.18 The detergent-solubilized protein is mixed with solubilized lipids to form a homogenous solution of mixed protein-detergent and lipid-detergent micelles. Detergent removal then results in the formation of protein aggregates in the worst case, and in the progressive formation of proteoliposomes with large 2-D crystalline regions in the best case. Reconstitution begins once the detergent concentration reaches the critical micellar concentration (CMC). The respective affinities between the components of the ternary mixture dictate the progress of the reconstitution process. Ideally, a starting condition should be established where mixed detergent-protein and mixed detergent-lipid micelles have exchanged their constituents to the extent that the mixture consists mainly of ternary detergentprotein-lipid micelles. Assuming that the protein remains in its native, properly folded state during the solubilization and isolation steps, this ideal situation is likely to foster perfect reconstitution and possibly 2-D crystallization of a functional membrane protein. In the course of many 2-D crystallization experiments the rate of the detergent removal emerged as an important parameter that requires optimization: a slow rate allows the proteins to pack regularly, but exposes them for a longer time to the detergent, whereas disordered

proteoliposomes form when the ternary system is forced to reach the CMC too quickly. The third method concerns the reconstitution of the membrane proteins at the water-air interface by attaching the solubilized membrane protein to an active lipid monolayer prior to detergent removal.19 In this process, membrane proteins are concentrated at the monolayer, brought into a planar configuration and finally squeezed together during detergent removal. This approach is useful for membrane proteins that are present in small amounts and are stably solubilized only in low CMC detergents. All methods summarized in Figure 1 have in common that the detergent is brought below its CMC to foster assembly of a bilayer, into which the membrane protein should integrate. The methods generally used to bring the detergent concentration below the CMC include dialysis,18 adsorption of the detergent to Bio-Beads20 or to cyclodextrins,21 and dilution of the ternary mixture.22 In all methods the amount of interspersed lipid must be minimized to ensure regular interactions between the membrane proteins. The pertinent interactions depend on the shape and surface charges of the components. For a given protein, the lipid-detergent mixture, pH, counter ions present and temperature must all be optimized. In addition, the concentration, the ratio of the respective components and the detergent removal rate are critical. This gives a multidimensional parameter space that needs to be experimentally sampled, a similar task to that carried out in 3D crystallization screens. The difficulty of such experiments is the management of the screens and the assessment of results. With 2-D crystallization the latter is particularly cumbersome because 2-D crystals cannot be detected by light microscopy, and screening by electron microscopy is time consuming. Precipitants are not generally required for 2-D crystallization; rather the most important factors appear to be the stability of the protein itself, the choice of lipid species, the density of the protein within the bilayer, and the surface charge (as controlled by pH and lipid head groups). There are three predominant morphologies adopted by the resulting 2-D crystals: i. flattened lipid vesicles with two, overlapping 2-D lattices; ii. tubular vesicles which retain a cylindrical shape and contain a helically organized array of membrane proteins; and iii. flat sheets composed of a single or double-layered coherent array of lipid embedded proteins. These 2-D crystals are very thin (50–80 A˚ for single-layered crystals, 100–160 A˚ for double-layered crystals and some 100 A˚ for tubular crystals) and are thus ideal samples for highresolution 3-D cryo-EM.

5.6.2.2

Sample Preparation

For electron microscopy, samples need to be either dehydrated or quick-frozen and kept at temperatures close to liquid nitrogen for transfer to the vacuum of an electron optical system. Dehydration not only changes the native environment of biological samples, but also exposes them to surface tension forces. Embedding aqueous suspensions of macromolecular

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

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Detergent Lipid Protein (with his-tag) Micelle Biobead cyclodextrin Monolayerlipid

(a)

(b)

(c)

(d)

Figure 1 2-D crystallization methods. All methods are based on the principle to bring the detergent concentration in the aqueous phase below the CMC, forcing the detergent in the mixed micelles to partition into the aqueous phase. As result, mixed micelles merge to form larger structures and ultimately 2-D crystals. (a) Dialysis can be used to remove the detergent provided its CMC is 41 mM. (b) Bio-Beads or cyclodextrins adsorb detergent molecules and can be used for all detergents. Bio-Bead/cyclodextrin driven 2-D crystallization is particularly successful with low CMC detergents. (c) Dilution is a well-known method for functional reconstitution of membrane proteins. In spite of reducing the protein concentration it is also suitable for 2-D crystallization, because the protein is highly concentrated after integration in the bilayer. (d) The monolayer technique combines the Bio-Bead method with crystallization at the air-water interface. This method works only with low CMC detergents because of the necessity to preserve the lipid monolayer. The latter incorporates special lipids having a high affinity for the solubilized protein (e.g., by recognition of a specific tag). By courtesy of Thomas Brown.

complexes or membrane fragments in a heavy metal salt solution provides support against surface-tension-induced compression of the biological structure during dehydration, and it produces a high contrast due to the strong scattering of the heavy atoms. However, heavy metal salt solutions create unphysiological ion strengths, and for uranyl salts a pH of around 4. Embedding the biological macromolecules in a sugar solution provides similar support under physiological conditions and maintains some hydration, yet the contrast produced by such samples is low, since sugar and protein have a rather similar electron scattering power. Hence this preparation method is mainly used for electron crystallography,5 because only ordered structure contributes to the crystallographic signal, not randomly arranged sugar molecules. To achieve atomic scale resolution, the flatness of the 2-D-crystals is a most important factor. Special molybdenum grids are utilized that produce a flat carbon film even after cooling the sample to the temperature of liquid nitrogen or helium.5

Charging the irradiated area can introduce image and sample instabilities and hence blurring of the high-resolution information. To overcome this problem, the double film sample preparation technique has been introduced, where the second carbon film reduces the instabilities.23

5.6.2.3

Image Formation

Modern electron optical systems comprise field emission electron guns operated at 100–300 kV that provide a highly coherent illumination and a wavelength l of 0.04–0.02 A˚. Magnetic lenses with spherical aberrations around 2–4 mm shape the illuminating beam, and collect the electrons scattered by the object to form an image at typically 50–100 000 fold magnification. Owing to the short wavelengths, atomic scale resolution can be reached at a small collection angle, YE0.61 l/d, hence electron optical systems exhibit a large

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Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

300 kV

Thickness (nm)

T D



0° 45° 60°

1000



d

100

0

10

–2 10 1

(b)

(a)

2

5 10 Resolution (Å)

5

3.4

2.5

Å

(c)

Figure 2 Image formation in EM. (a) The diffraction limit dictates the ultimate resolution of an optical system: d ¼ 0.61 l/Y. The depth of focus D corresponds to the distance between locations where the diffraction disc and the geometric discs are equal. Scattering centers within the resolution volume shaded in gray cannot be resolved (see Eq. 1). An optical system that collects waves emanating from scattering centers of a tilted 3-D object whose projected thickness T/cos(a)oD will produce the true 2-D projection of the 3-D object in the image plane. (b) Log-log plot of the resolution limit against the depth of focus D, as given by eqn [1]. The ordinate displays the maximal thickness T of the sample, which would be within the depth of focus D for a given resolution and a given tilt angle. For example, at a tilt of 601, a resolution of 5 A˚ is achievable at 300 kV acceleration voltage with a sample thinner than 200 nm. If a resolution of 20 A˚ is to be reached, the sample slab can be even 1.5 mm thick (indicated by the dotted lines). Adapted from Philippsen, A.; Engel, H. A.; Engel, A. The contrast-imaging function for tilted specimens. Ultramicroscopy 2007, 107(2–3), 202–212. (c) The contrast transfer function (CTF) of a 200 kV field emission transmission electron microscope whose beam divergence is 0.17 mrad. At Scherzer focus (blue), information is transferred without phase reversal and significant strength from 20 A˚ to 3 A˚. At 1 mm underfocus (black), information is transferred from about 40 A˚ up to a resolution of 3.5 A˚, albeit with reduced contrast at resolutions better than 5 A˚, resulting from the envelope function that reflects the partial coherence of the illumination. The phase reversal introduced by the CTF needs to be corrected computationally a posteriori. The spatial frequency is the inverse of the resolution labels given in A˚.

depth of focus (Figures 2(a) and 2(b)), as indicated by eqn [1]: Drd2 =ð0:61lÞ

½1

Here D is the depth of focus and d the resolution to be achieved. In addition, it is assumed that diffraction limited resolution can be reached after CTF correction (see below). Electrons interact strongly with matter, making it possible to depict thin unstained objects such as 2-D protein crystals, viruses, or small cells. Electrons are elastically scattered by the atom’s nuclei, whose mass is orders of magnitude larger than that of the moving electrons. Electrons are inelastically scattered by the inner and outer shell electrons, to which they transmit a fraction of their kinetic energy. While elastic electrons contribute to the coherent axial bright-field image that carries the high-resolution information on the 3-D arrangement of the sample atoms, inelastic electrons carry interesting chemical information. However, inelastic scattering is directly related to the beam induced specimen damage. Since only the elastically scattered electrons contribute to a high-resolution image, the coherent phase contrast imageformation is considered. A thin object that comprises only light elements and whose thickness is within the depth of focus (eqn [1]), is approximately described as weak phase object: tðx; yÞ ¼ 1 þ ifðx; yÞ;

fðx; yÞop=4:

½2

The function t(x,y) represents the two-dimensional (2-D) projection of the 3-D object. The amplitude distribution in the image plane is the coherent superposition of the unscattered wave and the elastically scattered waves. For an optical system

whose point spread function (psf) h(x,y) ¼ hr(x,y) þ i hi(x,y) is space-invariant, this superposition is described as the convolution of t(x,y) with h(x,y): aðx; yÞ ¼ ðhr ðx; yÞ þ ihi ðx; yÞÞ#ð1 þ ifðx; yÞÞ

½3

The intensity 9a(x,y)92 is recorded using film or CCD camera. Neglecting quadratic terms the image is: jaðx; yÞj2 ¼ 1  2hi ðx; yÞ#fðx; yÞ

½4

The imaginary part hi(x,y) is described by the inverse Fourier transform, FT1, of the phase contrast transfer function (CTF):24 hi ðx; yÞ ¼ FT 1 ½AðpÞsinfpðCs l3 p4 =2 þ Df lp2 Þg

½5

where A(p) describes the envelope of the CTF, Cs is the spherical aberration constant, Df the defocus, l the electron wavelength (about 0.02 A˚ for 300 kV electrons), and p the distance from the origin in the reciprocal space, and astigmatism is assumed to be negligible. The CTF for weak phase objects is displayed in Figure 2(c). Since the electron optical system introduces a phase difference of p/2 between the scattered and unscattered electrons, the axial bright-field mode corresponds to the Zernike phase contrast mode of light microscopy. The contrast is weak when the microscope is operated close to focus because the prominent low-resolution features of the specimen are transferred with small amplitude (Figure 2(c), CTF in blue). However, the contrast can be enhanced by moving out of focus, since frequency bands exhibiting a transfer coefficient 40.5 move towards lower resolution (Figure 2(c), CTF in black). Alternatively, electron

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

optical phase plates akin to the Zernike phase plate have been explored recently.25 In any case, the phase shift introduced by the electron optical system has to be corrected to facilitate the image interpretation (see below). The phase shift of electrons scattered elastically by an atom is proportional to the coulomb-potential of this atom. Therefore, the ensemble of all electrons singly scattered by a specimen produce a projection of its coulomb potential, which is dominated by the atom’s nuclei rather than the electron shells as in X-ray crystallography. It should be noted that this simplification is related to the large depth of focus provided by high-voltage transmission electron microscopes, as indicated by eqn [1] and illustrated in Figure 2. Figure 3(b) shows the power spectrum of a 2-D crystal image. This spectrum reveals: i. discrete spots representing the crystal information (see below); ii. concentric rings (Thon-rings) with gaps in between (the zero-crossings of the CTF, see Figure 2(c)), where no structural information is available; and iii. the envelope modulation decreasing the signal intensity at higher resolutions. Thon rings and envelope function reflect the specific nature of the CTF. The decrease of contrast towards high resolution (envelope function) results from the partial coherence of the electron beam. Modern electron microscopes are equipped with field emission guns exhibiting a high degree of coherence, so that the contrast decrease is acceptable even at high resolution. The Thon-rings with alternating positive and negative contrast are the result of the phase shift introduced by the objective lens. The great advantage of a modern field-emission electron microscope for directly acquiring the phase information out to atomic scale resolution is watered down by several experimental difficulties. First, the instrument has to be stable and installed in a field- and vibration-free environment, prerequisites that are routinely reached. Second, beam-induced damage not only changes the specimen structure, but leads to specimen charging. Since these charges induce specimen movements and act like an electrostatic lens image instabilities occur. If the sample plane is perpendicular to the optical axis, the overall effect is quite small: focus changes occurring during

97

image acquisition may have an influence only at very high resolution. In this case it is possible to measure the zerocrossings of the CTF accurately (see Figure 3(b)) and to correct the defects in the Fourier transform of the micrograph to retrieve both the phase and amplitude information. After this CTF-correction, the electron microscope performs close to a diffraction-limited optical system. However, when the sample is tilted for collecting the 3-D information (see below), the electrostatic lens which builds up during irradiation introduces an image shift, a problem that can be solved satisfactorily by using the double film technique.23 Third, a further optical defect is related to the changing focus for tilted specimens: the point-spread function is not space invariant, and the commonly used CTF correction is only an approximate measure to eliminate the phase distortions of the electron optical system.26,27

5.6.2.4

Electron Diffraction

When a parallel beam irradiates a crystal and the diffracted beams are brought to focus in the diffraction plane by the optical system, the resulting pattern can be recorded by a CCD camera, whose dynamic range is far better than that of photographic film. Electron diffraction is neither affected by the CTF nor by specimen movements. Moreover, the depth of focus is even larger than in case of imaging, since a small spread of diffraction spots can be tolerated. Therefore, electron diffraction is much more effective for the collection of highresolution information than imaging, although the phase information is not retrieved (Figure 3(c)). The directly measured amplitudes can be combined with the phases from the images during image processing. Electron diffraction is not an absolute requirement for determining a structure, but it allows a rapid judgment of the crystal quality, helps in correcting the CTF, and provides suitable high-resolution information for molecular replacement methods.6 To exploit the capacity of the electron microscope to acquire amplitude and phase information for crystallographic measurements, the primary prerequisite is the availability of highly ordered, thin crystals.5 They should exhibit lateral dimensions of several microns over which the crystallinity should be perfect.

Figure 3 Image, power spectrum and electron diffraction. (a) The electron micrograph of a large 2-D crystal does not reveal its crystallinity, but its homogeneity. The scale bar represents 0.5 mm. (b) The optical transform of such a micrograph shows sharp diffraction maxima and the effect of the CTF. The reflection indicated is at 9.1 A˚ resolution. (c) The electron diffraction pattern of a similar 2-D crystal exhibits diffraction maxima with a distinct four-fold symmetry. The reflection indicated is at 2 A˚ resolution. Unpublished data provided by Wanda Kukulski.

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5.6.2.5

Data Acquisition

Beam damage induced by inelastic interactions of impinging electrons with sample atoms dictates the maximum electron dose a 2-D crystal may take before discernable structural changes occur. This dose depends on the sample temperature: typically recording doses of 20 electrons/A˚2 can be applied if the sample is kept below 10 K.5 Hence, not only the best possible electron optical system is required, but the sample should be cooled to the temperature of liquid helium or at least to that of liquid nitrogen. 2-D crystals suspended in sugar solutions are adsorbed onto flat thin carbon films, semi-dried and frozen or vitrified, and are transferred to the cold sample stage. Images are acquired after the low temperature is reached and the stage has equilibrated. To minimize beam damage, crystals of potential high quality are identified at low magnification by their uniform thickness and characteristic shape (Figure 3(a)). Often, the entire grid is rapidly scanned in this mode and positions of interest are stored. The microscope is then adjusted for recording the high-resolution data. Either diffraction patterns or images are recorded, the former with a high resolution CCD camera, the latter preferably with photographic film. In special cases it is advantageous to record both a diffraction pattern and then an image. This allows the respective diffraction pattern to be properly classified based on the phases retrieved from the image.28

5.6.2.6

Data Processing

Each image represents a projection of the crystal, or after Fourier transformation, a central section through the 3-D Fourier transform (3-DFT) of the crystal (see Figure 4). Since 2-D crystals are periodic in (x,y), but have a single unit cell thickness in (z), the molecular transform, a smooth complex 3-D function, is sampled along z on lattice lines (h,k). Amplitudes and phases are obtained from the 2-DFT of the images after correction of the CTF. Each diffraction pattern, however, represents the central section through the intensity 93-DFT9.2 As recently reviewed, data processing proceeds according to the following general scheme:29 i. The CTF correction of the image is performed first, after the image has been Fourier transformed. As documented in Figures 2 and 3, this is a critical step for retrieving the phase information out to high resolution, since a small error in the determination of the CTF will lead to a wrong correction at high spatial frequencies, where the CTF oscillates rapidly. Importantly, this step needs to be executed before unbending takes place (see Figure 5(j)). A complication arises with images of tilted samples, as are required for extracting the 3-D information. Since the psf h(x,y) is not space independent, the linear systems theory yielding eqn [3] cannot be applied. A practical method to correct the optical defects in this case applies the CTF correction on stripes of similar focus.30 ii. The lattice parameters are measured from the Fourier transform of an image (2-DFT), in fact from its intensity 92DFT9,2 or directly from the electron diffraction pattern. The lattice is fitted by a least square distance minimization to

3D Object z

2D Projection u,v 2D FT Central section h,k u*,v*

z*

3D Fourier transform

h,k Figure 4 3-D structure determination by electron crystallography. To obtain a 3-D reconstruction from an object that is arranged in a 2-D crystal, 2-D projections are recorded at different tilt angles. The crystal lattice seen in the projection is defined by u,v in real space. The Fourier transform of a projection represents a central section through the 3-D Fourier transform (3-DFT) of the object. Projections (images) are processed to extract amplitudes A(h,k) and F(h,k) of the crystal’s h,k reflections, which are found on a lattice u,v. As result of foreshortening distances perpendicular to the tilt axis t in the projection, the lattice vectors of a projection (or its Fourier  transform) from a tilted crystal change: ut,vtau 0 ,v0 . The tilt geometry belonging to a crystal’s projection can thus be calculated from the respective lattice vectors. This information is used to calculate the elevation z for all h,k reflections, yielding measurements of the object’s 3-DFT in the form of triplets (A(h,k,z), F(h,k,z)). Since a 2-D crystal is not periodic in the z direction, the 3-DFT of the object is sampled on the 2-D grid h,k, but continuously along z. Experimental data are thus arranged in lattice lines, indicated in red vertical lines in the cube representing the 3-DFT. Lattice lines are interpolated to sample the object’s Fourier transform on a cubic raster. Back-transformation of the combined data finally leads to the 3-D representation of the object in the real space. By courtesy of Thomas Braun.

the diffraction peaks identified. Algorithms for automated lattice indexing have been developed.31 The lattice parameters are required to calculate tilt axis and angle, and subsequently the z for each reflection measured (Figure 4). iii. To extract the phase and amplitude information from the 2-DFT, sharp diffraction peaks are required. Hence all the parts of a 2-D crystal that are disordered and would contribute to background must be masked away or corrected by unbending. The latter is achieved by determining the position of all unit cells from the cross-correlation function

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

(2)

(1)

(a)

99

(b)

(c)

(3)

(4) (5)

(f)

(e)

(6)

(d)

(9)

(7)

(h1) (8)

(h2) (h)

(g)

Δf=1m

(i)

Δf=2m

(j)

Figure 5 Optimized information extraction from an image of a GlpF 2-D crystal. The raw image (a) is Fourier transformed (step 1) and the crystal lattice is indexed in the power spectrum 92-DFT92 of the raw image (b). Note that in this case, two crystalline layers of the flattened crystalline vesicle have to be separated. For the Fourier peak-filtering, the diffraction peaks containing all the crystal information are transmitted with weight 1, while the signal outside the mask-area (containing the other crystal layer and noise) is set to 0 (step 2, c). The filtered image (step 3, d) reveals the packing of the crystal. To unbend the 2-D crystal, a reference (e) is selected from D (step 4) and a cross-correlation (step 5) with the raw image is calculated. The cross-correlation (f) shows the positions of the unit cells. These can be compared to the fitted lattice (step 6) and difference vectors can be generated (g). This information is used to unbend the crystal and to eliminate badly distorted regions (step 7). As result, the spots of the power spectrum become sharper (h): In inset h1 peak 5,3 (indicated with a circle) is shown before unbending and in h2 after unbending. Amplitudes and phases of the spots are read out from the 2-DFT of the unbent lattice and combined with the data of other crystals (step 8). The 3.7 A˚ map of GlpF (i) revealing the typical tetrameric structures of aquaporins show the result of this procedure. The map is used as a new reference for refining the data extraction (step 9). In (j) the point-spread functions for Df ¼ 1 mm and Df ¼ 2 mm are displayed. As indicated, disrupting the psf by unbending would corrupt the information in the image, and phase errors and noise are introduced. Hence CTF correction is the first step in processing the image of a 2-D crystal. All insets are magnified 4  . GlpF unit cell has a size of 104 A˚, and the psf panels are 300 A˚ wide. By courtesy of Thomas Braun.

with a first average unit cell projection obtained from the uncorrected 2-D crystal. The displacement vector field between the fitted lattice and the actual unit cell positions is an ideal indicator of crystal quality (Figure 5(g)). The

possibility of such corrections is the great advantage of recording an image of a 2-D crystal rather than a diffraction pattern. However, unbending needs to be applied with care. For instance, unbending of a perfect membrane

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Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

protein 2-D crystal that is adsorbed on a bent carbon film would induce errors, as averages of crystal regions that do not have the same tilt will subsequently be calculated. Different unbending procedures are detailed in.29 Theoretically, it is possible to extract atomic scale resolution from small or fragmented crystals that do not exhibit sufficiently large highly ordered areas for electron diffraction. Unbending the image of a 2-D crystal and measuring the amplitude and phase of every (now sharp) diffraction order provides the averaged unit cell, or the projection map of the particular 2-D crystal, under ideal conditions down to a resolution of 4 A˚. On the other hand, electron diffraction of large, highly ordered crystals can deliver information to a resolution of 3 A˚ or beyond (see Figure 3 and ref. 6). The intensity of all peaks is measured by integration over the extent of the peak and subtraction of the local background. The signal-to-noise ratio (SNR) obtained in this way is used to weight the contribution of the respective peak. iv. Data collected from many images or diffraction patterns need to be merged to populate the 3-DFT as shown in Figure 4. To this end the projection maps obtained in step (iii) need to be correctly centered. Errors in this step introduce errors in the phase F(z), which are usually identified during lattice line fitting. To obtain the amplitude A(z), correct scaling of the data sets from single images or diffraction patterns is important. Scaling is optimized during refinement and lattice line fitting. A common strategy for merging projection maps taken over a tilt angle range of 01–701 is to merge the data from lower tilt angles, and to subsequently add data from samples titled at higher angles. Once the merging is achieved, lattice line data are fitted by a continuous function, and sampled at regular distances Dz to calculate the 3-D potential map by inverse Fourier transformation. From this first 3-D map comprising the total data set, projection maps can be calculated along any projection direction for subsequent refinement cycles. The goal of this procedure is to improve the centering, the tilt parameters, and the scaling of the respective projection maps. For electron diffraction patterns, refinement cycles improve the tilt parameters and the scaling. In advanced refinement cycles it is also possible to eliminate projection maps or diffraction data that deviate strongly from the overall data set, and thus would introduce noise. Such data processing is critical to extract all the information initially transferred by the electron microscope to film or to a CCD camera. It is a laborious process that contributes to the slowness of electron crystallography. Efforts are currently invested in improving the automation of data processing as well as the accuracy of the critical algorithms involved.31–33 Automation of data acquisition and processing will contribute to making electron crystallography a more widely used method.

5.6.2.7 5.6.2.7.1

Examples of Structures Solved by E-crystallography Bacteriorhodopsin

The first membrane protein studied by electron crystallography, bacteriorhodopsin (bR), is a light-driven protein

pump assembled into a highly ordered 2-D crystal by nature.34 These ‘purple membranes’ gave birth to electron crystallography through the collaborative effort of Nigel Unwin and Richard Henderson at the Laboratory of Molecular Biology, MRC Cambridge.3 Meanwhile six atomic structures obtained by electron crystallography have been deposited (PDB entries 1FBB and 1FBK; 2AT9; 1AT9; 2BRD; 1BRD). Of particular interest was the first visualization of the annular lipids surrounding a membrane protein35 and the study of Subramaniam and Henderson on the movement of helices F and G during the photocycle,36 which provided insight into the proton pumping mechanism (Figure 6). Compared to this, 80 wt and mutant bR structures in different activation states have been produced after the first X-ray structure has been published in 1997,37 mostly of the full-length protein and all by X-ray crystallography with the exception of the single structure achieved by solid state NMR.38 This situation is typical for progress in the structural determination of membrane proteins: Once the ice was broken by the invention of the cubic phase method for producing 3-D crystals of bR,39 X-ray crystallography with its powerful tools for acquiring diffraction data and software for solving structures simply took over. Yet, it was the pioneering work at the MRC Cambridge that paved the way towards a deep understanding of the bR photo cycle.

5.6.2.7.2

Aquaporins

Aquaporins (AQPs) represent an ancient family of small (E30 kDa) ubiquitous membrane channels that facilitate the permeation of water (aquaporins) and small, uncharged solutes (aquaglyceroporins). These proteins are present in all kingdoms of life, indicating their central role in maintaining the normal physiology of all organisms. The first demonstration of a biological water channel was achieved by Peter Agre and coworkers, who expressed a 28-kDa membrane protein of red blood cells in Xenopus oocytes.40 This protein was subsequently named Aquaporin-1 (AQP1). Electron crystallography of human AQP1 delivered the first atomic structure of an AQP41 (Figure 7). The novel AQP fold thus discovered consists of two pseudo-symmetrical halves, each comprised of three transmembrane a-helices and a reentrant loop, inserted into the membrane in opposite orientations and connected by the extended extracellular loop C. The six transmembrane helices H1 to H6 form the periphery of the monomer and surround the two re-entrant loops. Re-entrant loops B (between helices H2 and H3) and E (between helices H5 and H6) enter from opposite sides of the membrane and connect with each other in the center of the membrane through the proline residues of the two highly conserved NPA motifs. After interacting with each other, the re-entrant loops turn back and form the two short pore helices HB and HE. The structure also provided first insights into the water specificity and proton exclusion mechanisms of the protein. In particular, the structure revealed the importance of the two NPA motifs, both for stabilizing the AQP fold as well as for proton exclusion.41 Importantly, AQP1 was demonstrated to be fully active when reconstituted into highly ordered 2-D crystals long before its structure was determined.42 Shortly after publication of the AQP1 structure, X-ray crystallography made the structure of GlpF, the bacterial aquaglyceroporin, available.43 Another EM

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

D

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Figure 6 Conformational changes in bacteriorhodopsin during the photocycle as determined by electron crystallography to 3.2 A˚ in-plane and 3.6 A˚ vertical resolution.36 The unilluminated native bR (PDB 1FBB) is shown in blue, whereas the activated D96G, F171C, F219L triple mutant is in gray (PDB 1FBK). The native retinal is depicted in yellow. (a) Helix F rotates clockwise to open the retinal-containing pocket to protonated water molecules on the cytoplasmic side of the membrane. (b) The view from the cytosolic side into the retinal-containing cleft illustrates how water can access this pocket after bR activation. Scale bar: 10 A˚. The 3-D difference density map between the activated triple mutant and unilluminated native bR has been calculated from electron diffraction data and images. (c) Section of the difference density map close to the cytoplasmic boundary where the largest structural differences are observed in the map. The view is from the cytoplasmic side along an axis perpendicular to the plane of the membrane. Difference densities were contoured at 3s (blue, positive densities; green, negative densities). Prominent differences are in the vicinity of helices F and G. (d) Section close to extracellular boundary. The main change in this region is a positive feature localized to the vicinity of Arg 82. Panels (a) and (b) were produced with CHIMERA, while (c) and (d) were adapted from Subramaniam, S.; Henderson, R. Molecular mechanism of vectorial proton translocation by bacteriorhodopsin. Nature 2000, 406, 653–657. Copyright by Nature.

Extracellular

Loop A

Loop C H2

*

*

HE

H5

H3

H4 H6

*

*

HB

H1 –COOH

C-terminal NH2– helix (a)

(b)

Cytoplasmic

Figure 7 AQP1. (a) Aquaporin-1 as all other aquaporins and aquaglyceroporins exist as tetramer, which houses four independent pores (). As result of a gene duplication, the monomer consists of two similar halves that are structurally related by a quasi two-fold symmetry with the symmetry axis running through the center of and parallel to the membrane (red line). (b) The color coded AQP1 monomer reveals the AQP fold, with H1 (dark blue), H2 (light blue), HB (blue-green), H3 (green) forming the first half of the protein and H4 (yellow-green), H5 (yellow), HE (dark yellow), and H6 (orange) the second half. Important are the two half-helices HB and HE that emanate from the center outwards, as they are key to the pore structure. AQP1 exhibits a prominent loop A that connects H1 and H2, and an unstructured long loop C connecting the first and second part of the protein. The short C-terminal helix (red) was resolved by X-ray analyses.45 Adapted from Walz, T.; Fujiyoshi, Y.; Engel, A. The AQP structure and functional implications. In Handbook of Experimental Pharmacology; Beitz, E., Ed.; Springer-Verlag: Berlin, 2009; Vol. 190, pp 31–56. Copyright by Springer.

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Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

structure of human AQP1,44 and the X-ray structure of bovine AQP145 followed. Since then many aquaporin structures have been solved by X-ray and electron crystallography, and they demonstrate the common structural features shared by all aquaporins.46 AQP0 is the most abundant membrane protein in lens fiber cells and was initially named Major Intrinsic Protein (MIP).47 Like AQP1, it forms a channel that is highly specific for water, but it also mediates the formation of membrane junctions and was initially even thought to be part of gap junctions. The adhesive properties of AQP0 were first demonstrated in vitro by showing that reconstituting AQP0 into proteoliposomes induced the vesicles to cluster.48 Later studies established that proteolytic cleavage of its cytoplasmic termini increases the adhesiveness of the extracellular surface of AQP0.49 Electron crystallography of such double-layered 2-D crystals, grown with a mixture of full-length and cleaved AQP0 purified from the core of sheep lenses, eventually yielded a density map at 3 A˚ resolution, which made it possible to build an atomic model for the AQP0-mediated membrane junction.50 The atomic model identified the junction-forming interactions and also suggested that the water channel in junctional AQP0 may be in a closed or low-conductivity state. By subsequently solving the AQP0 structure to 1.9 A˚ resolution, electron crystallography not only showed its capability to reach very high resolution, but in addition revealed how lipids interact with a membrane protein. Nine lipid molecules surrounding each AQP0 subunit were resolved, making it possible to describe the interaction of AQP0 with its annular lipids6 (Figures 8(c) and 8(d)). Comparison with the X-ray structure of AQP0 in a detergent micelle demonstrated the stabilizing effect of lipids on the residues they contact.51 Most recently, a 2.5 A˚ resolution structure of AQP0 in a bilayer formed by E. coli polar lipids provided the first view of a membrane

1.8A

protein in two different lipid environments, and allowed the deduction of the first principles that govern the interaction of annular lipids with membrane proteins,52 demonstrating the importance of electron crystallography for assessing proteinlipid interactions. AQP4 is the predominant water channel in the mammalian brain, where it is expressed in glial cells.53 AQP4 forms orthogonal arrays in the plasma membrane of astrocytes and ependymocytes, which vary in size depending on the physiological conditions of the water homeostasis in the brain. AQP4 is expressed in two splicing variants, AQP4M1 and AQP4M23,54 of which only the shorter splicing variant AQP4M23 promotes the formation of large square arrays. The longer splicing variant AQP4M1 appears to restrict orthogonal array formation due to palmitoylation of N-terminal cysteine residues that are missing in AQP4M23.55 Electron crystallography produced a density map of rat AQP4M23 at 3.2 A˚ resolution, which made it possible to build an atomic model28 (Figure 9(a)) that was later confirmed by an X-ray crystal structure of human AQP4.56 AQP4M23 formed doublelayered 2-D crystals, and the interactions between AQP4 molecules in the adjoining membranes were mediated by a short 310 helix in extracellular loop C, suggesting that AQP4 may be involved in the formation of membrane junctions (Figures 9(a) and 9(b)). The notion that AQP4 makes membrane junctions was further supported by expression of AQP4 in L-cells, which resulted in cell clustering,28 and a recent, higher resolution, electron crystallographic structure of AQP4M23, which showed that AQP4 in one membrane interacts with lipids in the adjoining membrane.57 AQP4 thus became the second member of an AQP subfamily whose members are involved in the formation of membrane junctions. The 2.8 A˚ resolution structure obtained by electron crystallography of double-layered two-dimensional AQP457

PC7

Tyr 23

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PC1

PC3

Extracellular

2.0A 2.5A 3.0A

AQP0 H1 - Blue H2 - Light blue HB - Cyan H3 - Light green H4 - Green H5 - Yellow HE - Gold H6 - Orange CT - Red

PC9 Phe 144

5.0A

(b)

PC4 PC8 PC6

Cytoplasm PC5 Phitlt=100.13 Phi=100.23 Theta=7.82

(a)

(c)

(d)

Figure 8 AQP0. (a) Highly ordered double-layered 2-D crystals of AQP0 prepared by the carbon sandwich technique23 allowed electron diffraction patterns to be recorded exhibiting diffraction spots to a resolution beyond 2 A˚. (b) Two aromatic residues, Tyr 23 and Phe 144, that line the water pore in AQP0 are clearly resolved in the final 2Fo-Fc map of AQP0 refined to 1.9 A˚ resolution. (c) Vertical slab through the 2Fo-Fc density map with modeled lipid molecules, showing the two lipid bilayers in the double-layered AQP0 2-D crystal. (d) Nine lipid molecules that surround an AQP0 monomer in the 2-D crystal are resolved, with lipids PC1 to PC7 interacting with the protein, and lipids PC8 and PC9 belonging to the bulk lipids with no protein contacts. By courtesy of Thomas Walz.

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

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4.6 35.3

34.8 P139 P139

V142 V142

13.2

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5.9 11.6 (a)

(b)

(c)

Figure 9 AQP4. (a) Specific interaction at the extracellular surface of AQP4 tetramers dictate the packing into double-layered 2-D crystals, which has functional implications.28 While the side view shows that the AQP4 tetramers in the two membrane layers are staggered, the top view demonstrates that the entrances to the water pores in one membrane are partially obstructed by the four interacting tetramers in the other membrane. (b) Residues Pro139 and Val142 in extracellular loop C of AQP4 mediate the interactions between tetramers in the two adjoining membrane layers. (c) Water molecules in the AQP4 channel (red spheres) are clearly resolved in the 2Fo  Fc map contoured at 1.2s (marine mesh) as spherical densities.57 Distances (A˚) between water molecules and their closest protein atoms are depicted as dotted yellow lines. Distances (A˚) between neighboring water molecules are depicted as dotted yellow lines if in hydrogen-bonding distance, or as dotted cyan lines if the distance is too far for hydrogen bonding. A small density appeared in the channel, which has been interpreted as the quasi-stable position of a water molecule (marked by a white arrowhead and labeled as W3). The water molecules in the pore are designated W1–W8, and the temperature factors (in A˚2) of each water molecule are listed on the right side of the channel. By courtesy of Yoshi Fujiyoshi.

crystals further revealed eight water molecules in the channel, whose B-factors indicated different mobility (Figure 9(c)), shedding light on the water transport.

5.6.3 5.6.3.1

Atomic Force Microscopy Instrumentation

The principle of the AFM is simple: A sharp tip mounted at the end of a flexible cantilever is raster-scanned over a sample surface, while the cantilever deflection caused by the probe-sample interaction is measured via an optical system (Figures 10(a) and 10(b)). This signal drives a servo-system that moves either the sample or cantilever vertically to keep the latter’s deflection at a constant value. The surface topography is then reconstructed from this vertical movement. In this mode the probing tip always presses on the surface with a constant force during scanning (Figure 10(a1)).58 Alternatively, the AFM tip is oscillated rapidly in the vertical direction while scanning the sample. When the tip approaches the sample surface, a reduction in the oscillation amplitude is measured, and this is used to control the servo system (Figure 10(a2)). The tip oscillation reduces frictional forces, thereby minimizing damage and displacement of the

sample,59 so this mode is frequently used to image the surface topography of weakly immobilized biomolecules (e.g. single proteins and fibrillar structures). An even more sensitive variant of the oscillating mode is the dynamic mode AFM, where frequency shifts of the oscillating cantilever resulting from tipsample interactions are measured and used as feedback signal for the servo system (Figure 10(a3)).60 This mode is also called frequency modulation (FM)-AFM. The major advantage of all these imaging modes is that they can be executed in fluids, thus allowing biological macromolecules to be observed at work. Progress in instrumentation concerns AFMs that provide high scan speed,61,62 higher force sensitivity,60 and simultaneous acquisition of multiple signals.63 Operation in fluids imposes limitations and brings about technical difficulties. First, cantilevers that exhibit a Q-factor of about 104 in a vacuum and some 102 in air, have a Q-factor of o10 when operated in solution.64 Since the force sensitivity depends on the Q-factor, such a decrease is critical. Second, the resonance frequency of a cantilever drops by a factor of about two in liquid from its value in vacuum. Hence, the scan speed must be correspondingly reduced for scanning in liquid. Third, a conducting cantilever needs to be properly insulated for measurements of chemical reaction in solution, which requires complex microfabrication procedures.

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

(a1)

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Figure 10 Atomic force microscope (AFM) instrumentation and conditions for high-resolution AFM imaging in aqueous solutions. (a) Operation modes: (a1) Contact mode using the cantilever deflection as feedback signal; (a2) Oscillation mode using the oscillation amplitude as feedback signal; (a3) Oscillation mode monitoring the phase difference between driving force and cantilever oscillation. (b) Key elements of an AFM. A computer is used to control the movements of the sample and/or cantilever and store the surface contours measured. The instrument is operated with the sample adsorbed on a solid support (e.g., mica) in a buffer solution under ambient conditions. The bottom Figure shows a scanning electron microscopy image of a cantilever and pyramidal tip. Geometry and material of the cantilever dictates its mechanical properties (see eqns [6],[7]). The pyramidal tip has a spherical end with a radius as small as 10 nm, but tip surfaces are not smooth. (c) Small asperities of the AFM stylus contour the sample topography at high resolution. Interactions between AFM stylus and protein membrane can be divided in long- and short-range interaction forces. Long-range electrostatic repulsion forms a cushion that compensates forces applied to the tip. (d) The resolution of the contour measured is determined by the size of the asperity. (e) Force–distance curves recorded reveal electrostatic repulsion most clearly from (1) to (2). Increasing ion concentration and valency screens electrostatic interactions, revealing the van der Waals attraction at high ionic strength (lowest curve). Balancing van der Waals attraction and electrostatic repulsion is used to promote adsorption of the sample onto the support, as well as to minimize the forces between asperities of the AFM stylus and the protein. Adapted from Mu¨ller, D. J.; Fotiadis, D.; Scheuring, S.; Mu¨ller, S. A.; Engel, A. Electrostatically balanced subnanometer imaging of biological specimens by atomic force microscopy. Biophys. J. 1999, 76, 1101–1111 and Mu¨ller, D. J.; Engel, A. Atomic force microscopy and spectroscopy of native membrane proteins. Nat. Protoc. 2007, 2(9), 2191–2197 (copyright by Nature).

In spite of these issues, several successful designs of fast AFMs have been reported and encouraging results published. Fast scanning AFMs recording up to 1300 topographs within a second may in future help to capture the fast dynamics of membrane proteins.61,62,65 Moreover, progress in scanning electrochemical microscopy (SECM)63,66 suggests that electronic activation and direct observation of biological processes at the molecular level is feasible. These developments are closely linked to improved cantilever fabrication; small cantilevers are key to fast scanning, while insulated cantilevers with metal tips are needed for high-resolution scanning electrochemical microscopy. Indeed, spatial and time resolution are dictated by the cantilever and the sample’s physical properties. The geometry (length l, width w, and thickness t and shape), the density (r), and the elasticity module (E) of the material determine the cantilever’s mechanical properties, generally expressed by the spring constant (kL) and the resonance frequency (f0) in vacuum: kL ¼ E  w  t 3 =ð4l3 Þ

½6

pffiffiffiffiffiffiffiffi f0 ¼ c  t  f E=rg=l2

½7

Here, c is a shape-dependent constant that amounts to c ¼ 0.164 for rectangular cantilevers. For imaging biological samples in contact mode, the spring constant of the cantilever should be between 0.01 N/m and 0.1 N/m. The resonance frequency for these cantilevers ranges from 5 kHz to 50 kHz in vacuum and is slightly lower in air. When operating the

cantilever in fluid, f0 is reduced by a factor of 2–5. For example, a conventional rectangular silicon nitride cantilever (E ¼ 129 GPa, r ¼ 3440 kg/m3 67) with l ¼ 100 mm, w ¼ 40 mm, and t ¼ 0.4 mm has a spring constant of kL ¼ 0.08 N/m and a resonance frequency in vacuum of f0 ¼ 40 kHz. Another parameter characterizing the cantilever is the quality factor Q, which is defined by the stored energy W0 and the total energy loss per oscillation cycle DW:67 Q ¼ 2pW0 =DW

½8

In high vacuum, Q is a true cantilever parameter, because the energy losses result from internal damping mechanisms.67 However, in air or in fluid, the hydrodynamic damping by the surrounding medium dominates the energy losses.67,68 Although suitable contact mode cantilevers still have a quality factor of Q ¼ 10 to 100 in air, typical values drop down to Q ¼ 1 to 5 when they are operated in fluid. The resonance frequency and quality factor can be determined from the noise in the deflection signal,64 even when no oscillation mode is available. The thermal noise spectrum of the cantilever in water or air can be obtained by recording the deflection signal without a sample but in close proximity to the support and at a sampling rate that is at least twice the resonance frequency of the cantilever. For most applications, it is important to know the mechanical properties of the cantilever. Unfortunately, the microfabrication process is not accurate enough to warrant a cantilever thickness variation below 20%. As indicated by

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

eqn [6], this introduces large variations of the spring constant, making the routine characterization of cantilevers an imperative exercise. The spring constant can be calculated from the amplitude of the thermal deflection noise using the fluctuation method.64 To this end, the deflection signal (usually a voltage) needs first to be calibrated simply by pushing the cantilever upwards by a known amount and measuring the deflection voltage. Alternatively, the spring constant can be derived from the resonance frequency and quality factor of the cantilever, together with its top view dimensions.68 Interestingly, calibration of the deflection signal is not required here. The method assumes that the energy losses are dominated by the hydrodynamic damping, implying that the density and viscosity of the surrounding medium (air or liquid) are known. This method is derived for rectangular cantilevers but can be extended to different shapes as well, if a reference cantilever is available.68 Both methods64,68 give an uncertainty of o20%. The minimal force (Fmin) that a cantilever can measure is ultimately limited by the thermal noise of the cantilever. This can be expressed as a function of the spring constant, resonance frequency, and quality factor: Fmin  Fth ¼ fð2BkB T=pÞðkL =f0 QÞg1=2

½9

where kB is the Boltzmann constant, T the absolute temperature, and B the measurement bandwidth.67 For a cantilever as mentioned above (kL ¼ 0.08 N/m and assuming f0 ¼10 kHz and Q ¼ 2 in fluid) that is operated at room temperature with a sampling rate equal to the resonance frequency (i.e., B ¼ 10 kHz), eqn [9] gives a minimal detectable force of about 10 pN. This equation helps to optimize cantilever dimensions to achieve high force sensitivity, within the boundary conditions determined by the spring constant (0.01–0.1 N/m) and the limitations of microfabrication. In addition, a high resonance frequency is desirable to allow high recording speeds for observing real-time dynamics. Both goals (i.e., a better force sensitivity and a higher measurement speed) can be achieved by reducing the cantilever dimensions.69 For example, a small rectangular cantilever with l ¼ 20 mm, w ¼ 5 mm, and t ¼ 0.16 mm has a 10 times higher resonance frequency in vacuum compared to the 100-mm-long, rectangular cantilever mentioned above, while exhibiting the same spring constant (0.08 N/m). Thus, in contact mode the sensitivity is enhanced by a factor of three for a small cantilever when operated at the same bandwidth B as a conventional cantilever or the measurement speed can be increased by a factor of 10 without information loss. Another main feature of the cantilever is the tip sharpness. Although suppliers specify tip radii not better than 10–50 nm, topographs of flat biological surfaces that exhibit a resolution better than 10 A˚ have been acquired routinely.9–11,70–72 Provided that long-range interactions (e.g., electrostatic forces) are compensated for, only the short-range forces determine the tip-sample interaction. Consequently, only the tip atoms nearest to the sample determine the lateral resolution, which is a considerably smaller area than would be expected based on the tip radius given by the manufacturer. The tips often have a small asperity that protrudes sufficiently to contour the finest surface structures. Such a small asperity can exert a

105

prohibitively high pressure on the underlying structure, inducing its deformation. Therefore, pH and ion strength are adjusted to balance the tip-sample interactions.71

5.6.3.2

Sample Preparation

Sample preparation methods for raster-scanning samples in buffer solutions all have the objective of attaching the specimen firmly to a support. Immobilization is required for the contactimaging mode, as even small lateral forces exerted by the tip tend to push away the structure to be imaged. Lateral forces are largely eliminated by imaging modes, where the tip is oscillated vertically, lifting it away from the sample surface most of the time. For small particles and filaments, the contact area between sample and support is small, hence such structures can be immobilized for contact mode imaging by chemical fixation.73 As result of large interaction areas, biological membranes are simply adsorbed to a chemically inert, hydrophilic, flat solid support. Adjusting the electrolyte concentration and pH of the buffer solution facilitates this: the pH dictates the surface charge of the membrane, while the electrolyte concentration determines the thickness of the Debye layer.74 Because most biological membranes have a negative surface charge, the repulsive electrostatic force resulting from the negative surface charge of mica or similar supports prevents adsorption unless the Debye layer is thin compared to the decay length of van der Waals forces. Such solid-supported membranes have allowed, and will still allow, important insights to be gained into the structure and function relationship of native membrane proteins. Solid supports that have proven suitable for high-resolution imaging of membrane proteins include mica,75 highly oriented pyrolytic graphite,66,76,77 molybdenum disulfide,66 template stripped gold,66,78 and template stripped platinum.66 Template stripped metal surfaces appear to be particularly useful for combined topographical and electronic measurements.66 Detailed, step-by-step protocols for preparing biological membranes and for AFM imaging have been provided.79,84 Although high-resolution imaging is possible exclusively on solid-supported membranes, certain question may not be addressed by this preparation method. For example, membrane proteins in membranes directly attached to the support often exhibit impaired mobility,80–82 because the gap between membrane and support is only 0.5–2 nm. Moreover, adsorption forces may influence the conformation of membrane proteins, and it is known that lipids of a solid-supported lipid bilayer can show different structural features to the lipids of a vesicle or a freestanding lipid bilayer.82 Various schemes have been proposed to circumvent this problem by using spacers that warrant a larger gap, or polymer cushions.79 Freestanding bacterial S-layers spanned over small wells have been imaged at high resolution by the AFM,83 representing an ideal situation. Nevertheless, there is room for further progress in sample preparation strategies for studying the structure and function of native membrane protein assemblies by AFM.

5.6.3.3

Data Acquisition

To achieve high resolution, topographs are recorded in buffer solution. In this case, mainly electrostatic and van der Waals

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Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

forces govern the tip-sample interactions. In biological systems measured in aqueous solution, van der Waals interactions do not depend on the ionic strength, decay rapidly, and are always attractive. Electrostatic interactions result from hydrophilic surfaces that are often charged in water. They are longranged and can be attractive or repulsive, depending on the sign of the surface charges, which, in turn, depend on the pH. Because the stylus (silicon nitride, Si3N4) is negatively charged at neutral pH, and biological membranes are often negatively charged as well, the electrostatic forces are in most instances repulsive. The surface charges can be screened with electrolytes, and the electrolyte concentration determines the decay of the electrostatic interaction. In this way, the electrostatic interactions can be controlled. The Derjaguin, Landau, Verwey, Overbeek (DLVO) theory describes these forces quantitatively and allows the interactions between a spherical tip and a planar sample to be modeled, providing clues to optimize the

recording conditions.71 As illustrated in Figure 10(c) the tip surfs on a cushion of electrostatic repulsion under optimal conditions, while the small asperity discussed above is in contact with the sample without inducing deformation.71 Figure 10(e) also demonstrates how the repulsive electrostatic forces between stylus and sample depend on the ionic strength. The strategy for optimizing the recording buffer has been detailed recently.84

5.6.3.4

Image Processing

After investing a significant effort to acquire quantitative topographic information of a native biological membrane at sub-nanometer resolution, it is worthwhile to scrutinize this information with digital image processing tools. The example shown in Figure 11 concerns the cytoplasmic surface of bR

Figure 11 Imaging of the purple membrane surface by AFM in its most native state. (a) Depending on the the force applied to the stylus the cytosolic surface structure of bR trimers changes. During the scan, the force was varied from E50 pN (top) to E100 pN (upper half), to E50 pN (center) and to E100 pN (lower half). (b) This transition from fully extended E-F loop (left) to the compressed or bent E-F loop is visualized by morphing averages of topographs recorded at different forces applied to the stylus. (c) In images of the extracllular bR trimer surface, structural details can be clearly distinguished.136 Many diffraction spots are visible beyond (5 A˚)1 (white circle) in the power spectrum as indicated by solid circles. Because many gray levels are resolved in this image of purple membrane, whose extracellular surface exhibits a corrugation of 6 A˚, the vertical resolution achieved in this case is better than 1 A˚. Image processing of topographs recorded with the AFM allows different properties of the molecular surface to be visualized: (e) Analysis of the low force state of the cytosolic bR surface. From left to right: Correlation average; standard deviation (SD) map; probability map of loop locations; related surface energy potential. The SD- and probability maps show that the E-F loop, which is the prominent protrusion at the periphery of the trimer, is rather flexible. Therefore, a large SD is observed at the site of the E-F loop, and a wide distribution of its possible positions is seen in the probability map. This translates into a shallow potential trough in the surface energy map. In contrast, loop A-B (arrowhead) is well localized, leading to a small signal in the SD map and a sharp peak in the probability map. (f) Analysis of the high force state of the cytosolic bR surface. From left to right: Correlation average; SD map; probability map of loop locations; related surface energy potential. The location of the E-F loop still reveals a high SD, yet a new stable feature is now unveiled in this region, the C-D loop (arrowhead). A central well localized signal in both conformational states is related to the presence of a lipid molecule at the three-fold axis. Scale bar represents 100 A˚, and the width of all maps in (e) and (f) is 77 A˚. The grey level ranges in the SD map in (e) corresponds to 0.1–0.5 A˚ and in (f) to 0.1–0.3 A˚. The vertical bar indicates 5 kT.86 All topographs were recorded at room temperature in buffer solution. Adapted from Engel, A.; Gaub, H. E. Structure and mechanics of membrane proteins. Annu. Rev. Biochem. 2008, 77, 127–148. Copyright by Annual Reviews.

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

(see below). Although all bR trimers are identical to the 2-D crystal displayed in Figure 11(a), their surface structure varies significantly, predominately at the periphery of the trimes. These changes are related to the flexibility of the bR surface rather than any noise introduced by the atomic force microscope, and the structural details of each trimer are still discernable. Interestingly, the variation of this region between different atomic models from X-ray crystallography is pronounced: the conformation of the flexible EF loop, which is involved in the contacts leading to the 3-D crystals, is dictated by the 3-D packing arrangement of the bR molecules.85 Image processing now allows a variety of crucial parameters to be extracted. To this end, the individual molecular images need to be aligned laterally and angularly. Although this is not required for highly ordered 2-D crystals, it can be accomplished to sub-nanometer accuracy with randomly oriented single particles as result of the high SNR of topographs recorded by AFM. From aligned single molecular topographs, the average and the related standard deviation (SD) of the height measurement are calculated pixel by pixel, yielding the maps displayed in Figures 11(e) and 11(f). The SD maps document that vertical measurements can achieve an accuracy of better than 1 A˚, provided that the sample is sufficiently stiff. Calculating two independent averages from all odd- and all even-numbered single particles then allows the resolution to be determined by comparing the Fourier coefficients at a given resolution of these two averages, yielding the phase residual and the spectral SNR as function of resolution.86 Both analyses indicate the lateral resolution to be 5 A˚. Vertical fluctuations will emerge from lateral displacements of protruding loops that connect the helices of bR. Therefore, it is advantageous to characterize variations in the trimer structure by calculating the probability p(x,y)d of finding a certain loop (or domain) at a certain position (x,y)d by mapping the corresponding peak positions of all individual bR trimers, as illustrated by Figures 11(e) and 11(f).86 This map is readily converted to a free energy landscape Gd using Boltzmann’s law: Gd ðx; yÞ ¼ kB T ln½pd ðx; yÞ

½10

where kBT is the thermal energy, which is 4.1 10–21 J at room temperature.

5.6.3.5 5.6.3.5.1

Insights from AFM Topographs Bacteriorhodopsin

When operating a commercial atomic force microscope under optimal recording conditions, the surfaces of the biomolecules are contoured routinely at a lateral resolution of better than 10 A˚ and a vertical resolution of around 0.1 nm. Purple membranes of Halobacterium halobium are ideal objects to demonstrate the capability of an AFM and to optimize the imaging conditions. These highly specialized membranes are native 2-D crystals assembled from lipids and bR, the archaeal proton pump.34 On AFM topographs bR is resolved as trimeric structures arranged in a trigonal lattice of 6272 A˚ sidelength.3 As result of the exceptionally high SNR of the AFM, structural details of single bR molecules can be observed, and the flexibility of the peripheral protrusions extending 872 A˚ above

107

the lipid surface and comprising a small number of amino acid residues can be visualized.11,86 To record such topographs, the force applied to the stylus was approximately 50 pN, preventing a force-induced conformational change of the loop connecting helices E and F. When the force increases towards 100 pN, these loops bend, thereby changing the conformation of the bR surface. Four distinct protrusions can then be recognized in almost every monomer and a further, smaller protrusion is present at the center of the trimers (Figure 11(b)). At B50 pN vertical force, the standard deviation of the height measurements was around 1 A˚ for most morphological features of the topography, but the EF loop exhibited a standard deviation of 2 A˚ (Figure 11(e)) reflecting its high flexibility.11 The calculated diffraction pattern of the topography in Figure 11(c), which shows the features of the extracellular surface, documents an isotropic resolution out to 5 A˚ and beyond.11

5.6.3.5.2

E. coli Outer Membrane Porin OmpF

An E. coli outer membrane contains approximately 105 porins to allow the passage of nutrients that are o600 Da in size.87 The bacterial outer membrane protein OmpF exhibits a b-barrel fold with short loops on the periplasmic and extended loops at the extracellular surface.88 Conductance measurements have shown that porin OmpF trimers exist in either open or closed states, depending on the transmembrane potential.89 The critical voltage above which channels close is Vc490 mV for OmpF, and depends on the ionic strength. As this voltage is larger than that expected for the outer membrane potential, the physiological relevance of voltage gating has been questioned. However, evidence that Vc is affected by pH has been reported,90 and that membrane-derived oligosaccharides, polycations, low ionic strength buffer, and pressure all lower Vc.91 Although the structures of several porins have been solved, the mechanism of channel closure was not understood. As suggested by Schulz,92 the narrowest part of the channel (referred to as the eyelet) with its particular distribution of positively and negatively charged residues is the favorite candidate for a voltage switch. A model of voltage gating based on oppositely charged moving domains has been discussed, indicating that the loop lining the eyelet is the moving part of the gate. However, experiments with engineered mutants, where the loop forming the eyelet was tethered by a disulfide bond, suggested that this hypothesis needed to be reconsidered.93 An early study of two-dimensional porin OmpF crystals with the AFM revealed two conformations of the extracellular surface,94 but the reason for the conformational switch was not identified. Porin membranes adsorbed to highly oriented pyrolytic graphite (HOPG), showed an amazing conformational change of the extracellular surface that has not been predicted by the atomic structure. This occurred when the electric field across the 2-D crystal was changed by applying a voltage to a platinum wire closely positioned above the membrane.76 At zero voltage, topographs of extracellular porin surfaces revealed the protruding domain triplets of 13 A˚ height (Figure 12(b)), but this aspect changed dramatically into doughnut-like structures of about 6 A˚ height when the voltage was increased to 4.5 V (HOPG negative; Figure 12(c)). Clearly, the voltage directly

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Figure 12 Voltage-induced conformational change of the extracellular surface of porin OmpF. (a) Comparison of the atomic model of a rectangular porin crystal with the raw topographs recorded in buffer solution, operating the AFM in contact mode at minimal tip-loading force. Such crystals often consisted of two layers with the extracellular surfaces facing each other.94 Topographs of the periplasmic OmpF surface displayed features that correlated directly with the periplasmic aspect of the atomic model rendered at 3 A˚. (b) To image the extracellular surface, the upper crystalline layer was removed with the stylus. Extracellular domains protruded 13(72) A˚ (n ¼ 78) from the lipid bilayer and revealed a substructure consistent with the extracellular aspect of the atomic model, notably a distinct cleft that was visible in the protruding domains. Crystals were adsorbed to HOPG and topographs taken in 10 mM Tris-HCl (pH 7.8), 50 mM KCl. The brightness range corresponds to 15 A˚. (c) The conformational change of porin OmpF induced by either low pH or a membrane potential is summarized by the morphed averages from extracellular surfaces of 2-D porin crystals recorded at different voltages (adjusted by the ion concentration gradient) or at different pH. (d) This conformational change can be modeled as a rotation of the extracellular domain about a hinge at the rim of the b-barrel. A cross-section of the monomer demonstrates the open conformation (left porin monomer), and the closed conformation (right monomer). The black arrow indicates the putative rotation of the extracellular domain, while grey lines indicate the membrane surface. By courtesy of Daniel Mu¨ller.

over the membrane was much smaller than the 4.5 V applied, but could not be estimated with sufficient accuracy. At intermediate voltages, overviews showed the transition from the triplet protrusions to the doughnut structures, the porin trimer being unambiguously identified by correlating the topograph with the extracellular surface derived from the atomic model (ellipses in Figure 12(b)). The transition between the two conformations was fully reversible. To demonstrate that this voltage-induced conformational change represented channel closure, 2-D porin crystals were adsorbed to mica in 0.3 M KCl (pH 7.8). Shielding the negative surface charges of porin and mica with a buffer containing 40.1 M KCl was required for adsorption of the sample through van der Waals attraction,95 which in turn led to an increased K þ concentration between the porin membrane and the mica. Flushing the liquid cell in the AFM with a low ionic strength solution (5 mM KCl, pH 7.8) created a 460-fold cation gradient. This corresponded to a Nernst potential of 4100 mV, which was expected to induce channel closure. If the channels remained open, ions between porin sheets and mica would rapidly equilibrate with the bulk solution, and the membranes would desorb as result of electrostatic repulsion between the porin membrane and the mica. Interestingly, most of the double-layered porin crystals desorbed immediately, whereas most of the single-layered porin sheets remained firmly attached for more than one hour, consistent with channel closure. Their extracellular surface revealed domains of 13 A˚ height in the high ionic strength buffer, but after exchanging it with that of low ionic strength

(Figure 12(c)) doughnut-like structures were observed that protruded 6(72) A˚ from the bilayer. These structures appeared similar to those seen at an applied membrane potential. Again, this conformational change was fully reversible. After flushing the sample with 0.3 M KCl and imaging the same patch, the extracellular domains were fully extended. Because porin gating was known to be sensitive to pH,90 the AFM was used to visualize any pH-induced conformation change of the OmpF surface. Between pH 4 and 3, the extracellular protrusions collapsed, converting from a single protrusion (Figure 12(d), left) into a bi-lobed domain (Figure 12(d), right), concomitant with a height change from 1372 A˚ at pH 7.8 to 671 A˚ at pH 3. Three such domains formed doughnut-like structures similar to those induced by a membrane potential or by an ion gradient. The reversibility of this structural rearrangement was demonstrated by imaging the same membrane patch after increasing the pH to 4 (Figure 12(c)); the extracellular domains snapped back to protrude from the membrane by 12(72) A˚. Indeed, between pH 4 and pH 9, and in 0.1–0.5 M KCl, loops forming the extracellular domains of the OmpF assume the conformation determined by X-ray analysis (Figure 12(b)88). But these domains were found by AFM to collapse upon applying a membrane potential 4Vc, or upon changing the pH to 3 (Figure 12(d)). In the latter case, the loops are likely to assume a different conformation, because their charge distribution changes profoundly upon lowering the pH. For steric reasons, and since the b-barrel trimer is a rigid structure stabilized by intermolecular interactions,88 the

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

conformational change summarized in Figure 12(d) suggests that the protruding domains probably move towards and collapse into the extracellular vestibule about the 3-fold axis. Such a conformational change would reduce the height of extracellular domains from 13 A˚ to 6 A˚ and close the channel entrance. Because under physiological conditions a voltage beyond Vc is unlikely to occur, the AFM results strongly indicate that it is the acidic pH, rather than a voltage step, which leads to channel closure for the bacterial outer membrane porin OmpF under physiological conditions. The pH-driven closure of maltoporin was subsequently demonstrated by black-lipid membrane experiments,96 while recently the structure of an open and a closed conformation of porin OmpG showed the involvement of extracellular loops,97 and this conformational change was demonstrated by AFM under physiological conditions.98

5.6.3.5.3

Native Membranes

The best way to study membrane proteins under native or near-native conditions is to prepare native membranes directly and observe them in buffer solution. The atomic force microscope is currently the only instrument that allows imaging under such conditions in combination with a resolution better than 20 A˚. Interesting biological information has emerged from such studies. The first example concerns the disk membranes of murine retina where the AFM has unveiled the native conformation of rhodopsin molecules, which are arranged in rows of dimers99 (Figure 13). The importance of this observation is remarkable; for a long time, cross-linking

109

studies, bioluminescence resonance energy transfer and pharmacokinetics have suggested that G-protein coupled receptors (GPCR) work as dimers.100,101 Because rhodopsin is the first GPCR with a known structure,102 AFM data have provided a solid basis to model a GPCR dimer (Figure 13, inset) and to discuss possible interaction with the cognate proteins, such as arrestin and the G-protein heterotrimer.103 Another example is the outer mitochondrial membrane that houses specific proteins, which facilitate metabolic coupling and signaling between the cytosol and mitochondria. This membrane must be tight, because stress-induced release of cytochrome c leads to apoptosis. The voltage-dependent anion channel (VDAC), a general diffusion pore with a diameter of 20–30 A˚, mediates a majority of the molecular traffic. These porins have a molecular mass of around 30 kDa per functional channel, and have been found in all eukaryotic organisms. For membrane potentials 4920 mV9, VDACs are in a state that has a reduced conductance. Electron microscopy has shown that these pores form lattices whose unit cells comprise six VDACs when native mitochondrial outer membranes were treated by phospholipase A2.16 NMR and X-ray analyses have produced several atomic structures of the single channel.104 However, only recently has it been possible to reveal these channels directly in the native outer membranes of mitochondria from potato105 and from yeast.106 In some membrane domains, VDACs were found to be packed at high density like bacterial outer membrane porins,106 whereas in other domains, VDACs were loosely packed, exhibiting single pores and oligomeric clusters comprising two, three, four, and six channels105 (Figure 14).

Figure 13 Rhodopsin in the native disc membrane. After adsorption of osmotically shocked disk membranes on mica, their topography was recorded by AFM in buffer solution (20 mM Tris-HCl, pH 7.8, 150 mM KCl, and 25 mM MgCl2). The overview shows part of a disk membrane adsorbed with its extracellular surface facing the mica surface. The line pattern in the overview image recorded in error signal mode indicates tight packing with rhodopsin. Dense lines are seen as rows of rhodopsin dimers in the circular inset recorded in contact mode at high magnification.99 The disk membrane has a diameter of about 1 mm, while the double rows represent a paracrystalline array with a slightly skewed unit cell (a ¼ 84 A˚, b ¼ 38 A˚, g ¼ 851).103 Inset top left: Model of the arrestin-rhodopsin dimer complex. The dimer packing arrangement was elucidated from the AFM data. The theoretical model reflects the interaction of one arrestin molecule with the rhodopsin dimer.103

110

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins 4Å

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Figure 14 AFM topographs of the voltage-dependent anion channel in outer mitochondrial membranes. (a) Topography of outer mitochondrial membrane (OMM) patches. Two different types of surface are evident: (1) the OMM surface and (2) the mica surface. Scale bar: 0.75 mm. (b) VDAC proteins arranged in hexagons are visualized by contact mode AFM. The selected VDAC hexagons marked by broken squares are displayed in the gallery at the bottom. Scale bar represents 75 nm and the frame size of the magnified particles in the gallery is 18 nm. (c) The surface structure and organization of single VDAC proteins in the native OMM are revealed by FM-AFM. Various oligomeric states of the VDAC are displayed in the gallery at the bottom: monomers (marked 1), dimers (2), trimers (3), tetramers (4) and hexamers (6). The scale bar represents 75 nm and the frame size of the boxes in the gallery is 21 nm. By courtesy of Bart Hoogenboom and Dimitrios Fotoiadis.

The advantage of FM-AFM compared to contact mode AFM is demonstrated by the images of the voltage-dependent anion channel in native potato mitochondrial membranes.105 Whereas in contact mode small oligomers were hard to observe and single VDAC channels were not found, monomers, dimers, trimers, tetramers and hexamers were unambiguously identified by FM-AFM. This observation suggests that FM-AFM induces smaller lateral forces, thereby making the observation of single channels embedded in the bilayer possible. Topographs acquired by AFM of native membranes from different photosynthetic bacteria have provided new information on the architecture of the photosynthetic apparatus (reviewed in107). These membranes have been imaged at sufficient resolution to visualize individual subunits. Such images have also revealed the structural changes of the photosynthetic machinery during chromatic adaptation of R. photometricum to high-light and low-light growth conditions.108 An insightful atomic model of the light-harvesting system has subsequently been established by using the surface topography acquired by AFM and the atomic structures of individual subunits.109

5.6.4 5.6.4.1

Single Molecule Force Spectroscopy Instrumentation and Methodology

A modern commercial atomic force microscope offers both a high spatial resolution and force sensitivity down to some 10 pN. To measure forces of single molecular interactions, molecules are tethered to a support and tip and forces are measured while the cantilever is retracted from the support. Soft cantilevers (ko0.05 N/m) are used to record forcedistance curves during such events as the unfolding of a protein. Although a variety of chemical surface activation

techniques have been used, large 2-D assemblies such as reconstituted or native membranes14 and bacterial S-layers13,110 are simply physisorbed to mica as for imaging. The AFM stylus is then attached to an individual protein of such an assembly by pressing the stylus down with a force of B1 nN for approximately 0.1–0.5 s, which induces denaturation of some protruding domain that is thus stuck to the silicon nitride tip. The finding that physisorbed molecules may withstand pulling forces of several hundred pN before they detach indicates that multiple local interactions based on van der Waals forces, charge interactions and hydrogen bonds stabilize this contact.111 When the nN force regime needs to be probed, the proteins must be covalently attached, preferably via known natural or site-specifically introduced reactive amino acids (e.g. a cystein), on a surface-exposed domain that allows specific attachment of the protein to a gold-coated stylus. This covalent attachment not only allows an extended force range to be explored with the consequence of a prolonged time window for the experiments, but it also enables precise and absolute length measurements to be made during the unfolding process.14 However, although the percentage of full-length traces has increased to 90% with goldcoated tips, the price of a reduced resolution in imaging and a drastically decreased lifetime of the tip needs to be taken into account.14 The dissociation of biomolecular bonds is a temperaturedriven, dynamic process. To probe it over different timescales, the acquisition of force-extension curves is achieved at different extension rates, a method that is referred to as dynamic force spectroscopy (DFS). Quantitative ensemble measurements can only provide macroscopic thermodynamical quantities (e.g., the free energy of complex formation and/or dissociation). From DFS measurements, however, the virtual energy landscape of a single molecular interaction can be mapped, giving a detailed insight into the reaction pathway of

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

111

single molecular processes. Thermodynamical models of the rupture of a single bond are described in.112,113 A liner increase of unbinding forces with the logarithm of the loading rate has been observed.114 A similar behavior has been demonstrated for the unfolding of proteins, at least in the limit of small forces. This is a direct consequence of an exponential decrease of the bond lifetime with an applied force, which can be explained by assuming that the dissociation barrier DE0 is linearly decreased by applying a force F:112

tip is very low. This is achieved at low concentration and using linkers that have a length that is comparable to the diameter of the AFM-tip (about 50 nm). Nevertheless, rupture event series may still be observed. Therefore, it is the last rupture event that should be considered to measure the single molecule rupture force F.

DE ¼ DE0  xF

For single molecule force spectroscopy (SMFS), native biological membranes or reconstituted densely packed, or crystalline protein-lipid layers are prepared by adsorption to mica at appropriate pH and ion strength as described above. Since mica is negatively charged (  0.0025 C/m2 at neutral pH) specific interactions with positively charged residues need to be considered. Nevertheless, the wealth of force-distance curves which have been acquired on different membranes adsorbed to mica demonstrate that such specific interactions have so far not been observed.

½11

and that the bond lifetime t(F) is given by the Arrhenius expression: tðFÞ ¼ t0 expðxF=kB TÞ

½12

where t0 is bond lifetime without applied force. The bond length x describes the geometry of the energy landscape. It is interpreted as the distance between the ground state and an energy barrier along the dissociation path (the reaction coordinate) projected onto the direction of the applied force. In principle, many reaction pathways exist, but the applied force is expected to select a specific reaction coordinate in the multidimensional energy landscape. Thus, the system will follow the reaction pathway corresponding to the lowest energy barriers. From eqns [11] and [12], the thermal off-rate koff(F) of a bond under an applied force can be written as: kof f ðFÞ ¼ Kof f expðF=F0 Þ

½13

where Koff is the natural thermal off-rate for dissociation and F0 is a force-scale factor (F0 ¼ kBT/x). Although the bond lifetime is a relevant parameter to describe the kinetics of bond rupture, the quantity directly measured in a typical AFM experiment is the unbinding force. Because the rupture of a bond is a stochastic process, the measured unbinding force follows a distribution whose width is mainly determined by the force-scale factor F0.114 In this case, the most probable unbinding force (F) depends on the applied loading rate r as: F  ¼ F0 ln½r=ðF0 kof f Þ

½14

DFS measurements can be performed with an unmodified commercial atomic force microscope, possibly using external data acquisition capabilities that enhance sensitivity and flexibility of the instrument. To achieve reproducible measurements, the spring constants of all cantilevers used must be calibrated as described above. Because the temperature is a crucial parameter in these experiments, it should be controlled, for example using a regulated Peltier element, which is in contact with the buffer solution.115 When approaching the tip to the surface, nonspecific attachments may occur even if the support has been passivated or is a pure polymer surface. Thus, unspecific interactions need to be minimized using linkers [e.g., poly(ethylene)glycol (PEG) linkers] that shift the region where unbinding takes place away from the surface. Finally, to quantify the most probable value for the unbinding force of a single complex, conditions must be chosen in which the probability that two or more complexes are attached to the

5.6.4.2

5.6.4.3

Sample Preparation

Data Acquisition

For the investigation of membrane proteins two strategies have proven to be useful. While imaging the membrane proteins at high resolution, an individual protein can be selected by halting the scan and attaching the selected protein to the tip by increasing the force beyond the adhesion threshold, typically in the range of 0.5–1 nN. Upon retracting the tip, a force-distance trace can often be recorded during the unfolding and extraction process. A subsequent image of the same area allows the site where a single membrane protein was extracted to be identified (see Figure 15). Although this protocol brings additional information on the unfolding process and the molecular interactions, it is a laborious and timeconsuming procedure.13,14,110 Alternatively, a suitable membrane patch is identified at low magnification and the tip is then positioned above this patch to acquire the force curve in a semi-automated manner (Figure 16).120 Where necessary, the AFM tip can be used as a nano-scalpel to remove aggregates or the upper layer of a collapsed vesicle, thus exposing suitable membrane areas for force spectroscopy. Force curves can then be acquired on a point grid with an edge length of 150–300 nm and a linear point density of 0.125 nm1 (Figure 16(a)). Membrane proteins are attached to the AFM tip by pushing the cantilever ten times onto the membrane at each grid point with a force of 0.5 nN for 0.1–0.6 s. Retracting the cantilever for 0.25 s with a velocity of 0.5–1 mm s1 then produces a force-distance (F-D) curve. All F-D curves (comprising e.g., 4096 data points) are saved no matter whether a protein became attached to the tip or not. For both protocols, the cantilever’s force constant needs to be measured, usually exploiting the thermal noise method.64 Moreover, the cantilever deflection detected with a positionsensitive photodiode (PSD) needs to be additionally calibrated by bringing the tip into contact with the mica and then moving the cantilever even closer to the support and measuring the PSD voltage over a deflection regime between where the force increased linearly with piezo movement (typically

112

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

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Figure 15 Visualization and unfolding of a single bacteriorhodopsin (bR) molecule. (a) A purple membrane with its cytosolic surface facing the stylus was imaged at high resolution. The scan was stopped (circle) and the stylus pressed down for 0.1 s to catch a bR molecule. (b) Unzipping a single bR molecule occurs upon retracting the stylus and produces a series of force peaks that can unambiguously be related to the seventransmembrane a-helical structure of bacteriorhodopsin. (c) The subsequently recorded high-resolution topograph reveals the vacancy left by the unfolded bR molecule. The contour indicates the shape of a bR trimer. From Oesterhelt, F.; Oesterhelt, D.; Pfeiffer, M.; Engel, A.; Gaub, H. E.; Mu¨ller, D. J. Unfolding pathways of individual bacteriorhodopsins. Science 2000, 288(5463), 143–6. Copyright by AAAS.

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Figure 16 Automated acquisition and analysis of force-curves. (a) The cantilever deflection image shows a single-layered membrane (marked 1), densely packed with AdiC proteins (dots on this surface type) and the mica surface (marked 2). The white box on the AdiC proteoliposome (1) shows the area (frame size: 200 nm), where F-D curves were recorded. The zoomed area contains 625 positions at which measurements (e.g., ten measurements per position) were performed. (b) Filtering with minimal tip-sample separation tssmin ¼ 95 nm and a minimal force of FSMFS,thr ¼ 3.5  sF resulted in the import of 190 out of 22,000 F-D curves. Arrow marks curves that had no sharp rise, indicating some contamination between tip and sample. (c) After elimination of F-D curves exhibiting contaminations and a length that was incompatible with the once expected for AdiC, and final manual corrections, the data set looks much cleaner. Scattering plots of aligned force spectra and histogram of contour length values for N-terminal unfolding of N-His6-AdiC in the presence of 10 mM L-arginine, n ¼ 193, panels (d), (e); 10 mM agmatine, n ¼ 197, panels (f), (g); 10 mM D-arginine, n ¼ 196, panels (h), (j). By courtesy of Patrick Bosshart and Patrick Frederix.

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

from 300 pN to 1 nN). This calibration allows the PSD voltage to be converted into a metric deflection, and finally into a force by subsequent multiplication by the cantilever’s force constant.

5.6.4.4

Only the part of the F-D curve acquired during retraction needs to be analyzed. F-D curves are shifted vertically so that the average force in the baseline, defined as the final 5% of the data points in the tail part of the spectrum, is zero. Tip-sample separation values (tss) are computed by subtracting the corrected metric cantilever deflection from the position of the z-piezo. Usually force spectra are flipped vertically, so that pulling force peaks faced upwards – attractive forces are positive in this representation. In a first step, all F-D curves need to be filtered to extract suitable unfolding events for further analysis. A few simple criteria need to be met: 1. The tail of the spectrum should be flat, that is, close to 0; 2. The length of the acquired F-D curve should correspond roughly to the number of residues, that is the last event exhibiting a force 450 pN needs to occur at a minimum distance that is compatible with the full extension of the assessed protein during unfolding, but shorter than a maximum distance to ensure that a single protein has been unfolded; 3. Unspecific adhesion forces need to vanish within a short interval to eliminate F-D curves where some contaminant inhibited direct interaction between tip and addressed protein. In a second step, F-D curves need to be inspected visually to identify a common ‘fingerprint’. Curves with atypical unfolding patterns can be removed. The progress in cleaning up F-D curves is illustrated in Figures 16(b) and 16(c). Typically 100–200 classifiable unfolding traces are then aligned with respect to the most prominent unfolding peak (Figures 16(d), 16(f) and 16(h)). The characteristic shape of peaks in the F-D curves has been approximated with different models (see ref. 8). Equation 15 gives the often-applied wormlike-chain (WLC) model, where x is the extension, p the persistence length and L the contour length of the unfolded polypeptide chain: Fðx; LÞ ¼ ðkB T=pÞð1=ð4  ð1  x=LÞ2 Þ þ x=L  1=4Þ

barriers on the opposite side of the pulling tip. A shift of the barrier location by (4.8 nm/(0.36 nm)1) aa E13 aa achieves the compensation.118

5.6.4.5

Data Processing

½15

Using the WLC model and assuming a fixed persistence length of p ¼ 0.4 nm,116,117 F-D curves can be transformed into F-L curves.15 The advantage of this transformation results from the fact that force measurements F(x) are mapped into the contour length space, where barrier positions are directly visible, independent of the actual rupture force that depends on fluctuations and external experimental parameters. Aligned F-L plots can then be integrated in a histogram, and peaks can be fitted by Gaussian curves to determine the respective barrier positions with an accuracy of a single residue (Figures 16(e), 16(g) and 16(j)). To map the barriers onto the measured or predicted structure of the membrane protein which has been obtained, the membrane thickness needs to be compensated for unfolding

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5.6.4.5.1

Insights from Single Molecule Force Spectroscopy (Examples) Visualizing the Site where a Membrane Protein was Unfolded

Advances in single molecule force spectroscopy applied to soluble proteins and cells tethered between a support and a cantilever119 have stimulated experiments on bacterial surface layers13,110 and on bR.14 As illustrated in Figure 15, these experiments have provided not only information on the unfolding forces, but also revealed the structural changes related to the extraction of a single membrane proteins using the AFM tip as nanotweezer.

5.6.4.5.2

Localizing Unfolding Barriers

During protein unfolding, the force exerted by the cantilever is transmitted along the backbone into the membrane protein. The latter is stabilized in its conformation by the interplay between local forces, which in their complexity are represented by the potential energy landscape. The retracting tip drags the protein in the 3N-dimensional energy landscape along a chosen direction and may encounter a barrier, which needs to be overcome to ensure that unfolding continues. Two contributions act together in this process; the external force, which is transmitted through the backbone of the already unfolded part of the protein and thermal fluctuations (see eqn [13]). Therefore, simply waiting increases the likelihood that thermal fluctuations will overcome the remaining barrier, and if in addition an external force is applied, the barrier will be overcome at lower forces, the slower an external force is ramped. The point where the barrier is overcome will thus depend not only on the energy landscape but also on the load rate r.112 Since the AFM allows the position of the tip to be controlled with Angstrom precision, the barrier position can in principle be assigned with single residue accuracy by mapping F(x) into the forcecontour (FC) space (Figure 16). However, because backbone elasticity and thermal fluctuations alter the apparent length of the unfolded protein under load, a small correction needs to be applied (see refs. 8, 12). An interesting case is the SMFS analysis of AdiC, an arginine/agmatine exchager,120 because the barriers have been mapped on the predicted secondary structure, before the atomic structure was solved (Figure 16).

5.6.4.5.3

Localizing a Ligand Binding Site

Energy barriers may not only be signatures of the intramolecular bonds, they may also be modulated by the interaction of ligands with the protein. The capability of the AFM to localize such barriers with the precision of a few amino acids opens an avenue for exploring the energy landscape of the protein for signatures of ligand interaction. The feasibility of this approach has been demonstrated for the first time for the sodium proton antiporter in its active and its inactive state.121 Figure 17 illustrates that an additional peak

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Figure 17 Mapping a ligand binding site. (a) AFM topograph of reconstituted and 2D crystallized proton/sodium antiporters (NhaA) from E. coli. The inset shows a correlation average of the topography, revealing structural details of the membrane protein. The topography was recorded in buffer solution (200 mM NaCl, 20 mM Tris–HCl, pH 7.8) and exhibits a full gray level corresponding to a vertical range of 3 nm. Scale bars: 25 nm, and 10 nm in the inset. (b) and (c) Superpositions of E20 aligned force-distance (F-D) curves that have been recorded upon mechanically unfolding of single NhaA molecules in their active (b) and inactive (c) state. (d) Single F-D curve showing a detailed spectrum of force peaks. Each force peak denotes an interaction that has been established by an unfolding intermediate. Fitting each force peak using the worm-like-chain (WLC) model (red curves) reveals the contour length of the unfolded and stretched polypeptide. This contour length allows each specific barrier, which established during unfolding of NhaA to be located. SMFS experiments were performed in 150 mM KCl and 50 mM NaCl. (e) Secondary structure of NhaA with each sphere locating an interaction barrier that has been detected by SMFS as shown in (d). The interaction at a contour length of aa 225 (measured starting from the C-terminus) is the result of ligand binding (Na þ -ion) to the antiporter. The inhibitor (2-aminoperimidine, AP) binds to the ligand-binding site and establishes similar interactions as the ligand. However, in addition the inhibitor engages in an interaction at helix IX to deactivate the antiporter. By courtesy of Daniel Mu¨ller.

appears at an extension of roughly 65 nm upon activation in neutral pH. This can be attributed to a novel barrier around aa 225, which is located in the middle of helix V. A functional inhibition by 40 mM 2-aminoperimidine, however, results in the formation of a new barrier at around aa 85, which is located in the loop between helix X and XI. The authors interpret this unexpected finding by a ligand-induced stabilization of the loop, which together with other parts of the protein may form the binding pocket. This pioneering experiment opens the path for more ligands and binding pockets to be identified in the long list of health-relevant membrane proteins for which no high-resolution structural information is available.

5.6.4.5.4

Unfolding from Different Ends

Membrane proteins like bR have a preferred orientation. The different sides of the membrane thus make different sides of the membrane protein easily accessible for single molecule force spectroscopy. The individual structural elements of the protein may thus be unfolded in reversed order. One might argue in the first instance that the unfolding barriers are approached simply from the ‘‘backside’’. However, it becomes clear in a more profound analysis that entirely different unfolding pathways are taken when the protein is unfolded from the N and the C terminal ends, respectively. Nevertheless certain barriers (e.g. hydrogen bonds or salt bridges), may be highly localized and therefore give rise to a peak in the

Atomic Force Microscopy and Electron Microscopy of Membrane Proteins

unbinding force when approached from either side. Kessler et al. have demonstrated this for bR, which they unfolded from the cytosolic and the extracellular side.122 For each branch of the unfolding trace they fitted an elasticity curve and localized the barrier. It is interesting to note that the peak heights of the unfolding forces and thus the apparent barrier height dropped with increasing unfolding of bR. This has been interpreted as the decreasing interaction with the local environment of the protein, since parts of the protein may have already been extracted. The two sets of traces recorded from the two sides thus contain complementary information. Surprisingly, Kessler and coworkers,122 and also others (see ref. 12), found barriers not only in or between those structural elements, which are known to be stiff (e.g. a-helical rods), but also in loops that are not well resolved in structural investigations. In certain cases, the barrier positions were found to coincide when probed from both sides. These barriers are of particular interest since they must be stabilized in both directions, upstream and downstream. In one case, the segment of the protein between this given position and the C-terminal had already been extracted, while in the other case the segment towards the N-terminal is still integrated in the membrane when the barrier is probed.

possible at the level of individual helices. The next step was then to allow the proteins to refold into the membrane, and to investigate this process as a paradigm for protein folding. This has been achieved with the sodium proton antiporter,123 and with bR.124 Figure 18(b) shows the protocol for such an experiment. First helices G and F were unfolded. Subsequently, helices E and D were unfolded, and the tip was then lowered again, allowing the protein to reassemble in the membrane. Figure 18(a) shows that at a certain point the protein pulled the cantilever down. Therefore, the protein exerted mechanical work against the cantilever (see green area under the curve in Figure 18(a)). Subsequent unfolding confirmed that helices E and D had fully refolded. However, the majority of the folding attempts did not result in a successful but rather an imperfect refolding, which has been attributed to both the rearrangement of the void in the membrane as well as inadequate drift stability over the prolonged time spans needed for this kind of experiment.124 Since such experiments provide both the work of unfolding and the work of refolding, they may become the foundation for future analyses of membrane protein folding.

5.6.5

Perspectives

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Figure 18 Unfolding and refolding of bacteriorhodopsin (bR). (a) Unfolding traces (red) and refolding traces (blue) of a bR molecule whose helices F and G had already been unfolded. (b) Schematic representation of the experiment. Helices E and D were extracted from the membrane (1–4). Subsequently the cantilever was lowered, and the pulling force was recorded during refolding of the two helices (5–8). The next unfolding cycle confirms the complete refolding of helices E and D (9–13). The mechanical work performed by the membrane is highlighted in green. By courtesy of Hermann Gaub.

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environment. As demonstrated in reconstituted 2-D crystals in reconstituted 2-D crystals,42 membrane proteins retain their biological activity in these experiments. Because the molecular packing in 2-D crystals is less constrained than in 3-D crystals, conformational changes can be accommodated. This property has been used to study the activated state of the visual pigment rhodopsin (reviewed in125). Following absorption of a photon, isomerisation of 11-cis retinal initiates the photoactivation process, resulting in an equilibrium between meta-I and meta-II states of the GPCR. Rhodopsin is photoactive in 3-D crystals, but diffraction deteriorates dramatically after illumination; therefore, it is not yet possible to study the late metarhodopsin intermediates using X-ray crystallography. The structural rearrangements of this intermediate state can, so far, only be studied by electron crystallography in the more physiological environment of a 2D crystal.126 Membrane proteins composing a 2-D lattice are fully accessible to the aqueous medium, thus providing an optimal binding surface for ligands. This property minimizes ‘‘low occupancy’’ issues when the crystals are soaked with specific ligands or inhibitors. This property has been exploited to study conformational changes of the multidrug transporter EmrE upon binding of ligands of varying sizes.127–129 A wealth of biochemical, cell biological and physiological observations indicate the critical role lipids play in membrane protein assembly and regulation. However, structural information has been hard to collect. Therefore, the recent development of assessing membrane protein-lipid interactions by electron crystallography is most important, and is likely to provide further insights into these processes.6,52,57 Clearly there is a lack of massive instrumental and computational developments in the field. Nevertheless, the constant efforts of a few laboratories have recently led to progress in the automation of 2-D-crystallization and evaluation of crystallization screens.130–134 Moreover, automated processing of crystal images and diffraction patterns is finally emerging,32,33 fostering hopes for an improved throughput in electron crystallography. The AFM has opened avenues for visualizing single membrane proteins embedded in the bilayer under physiological conditions at sufficient resolution to identify and manipulate single polypeptide loops connecting a-helices11 or b-strands.76 This instrument has also made it possible to unfold membrane proteins quantitatively to probe the forces that dictate the fold of the protein,122 and to locate ligand binding sites.121 As result of the high SNR that the AFM imaging modes provide, imaging native membranes has become feasible at a resolution that allows individual membrane proteins to be recognized, generating important insights into the function of rhodopsin, VDAC, and bacterial photosynthesis.99,105,107 Advancements in AFM instrumentation and sample preparation have opened yet another field; the quantitative assessment of forces between and within cells. Rapid progress has stimulated ideas for new measurements that aim at deciphering the complexity of cellular behavior.134 Since this instrument is still relatively young, it is expected to provide a wealth of new data on the structure and function of membrane proteins in the cellular context.

Acknowledgments The authors are indebted to Wanda Kukulski, Thomas Braun, Thomas Walz, Yoshi Fujiyoshi, Daniel Mu¨ller, Hermann Gaub, Dimitrios Fotiadis, Bart Hoogenboom, Patrick Bosshart, Patrick Frederix, and Simon Scheuring for inspiring discussions and for providing their results. This work was supported by the Swiss National Science Foundation (SNF grant No. 3100A0-108299), the National Center of Competence in Research in Structural Biology, the University of Basel, and the Maurice E. Mu¨ller Foundation of Switzerland. The used AFMfacility was built up with contributions of the Swiss University Conference and JPK-Instruments AG, Berlin, Germany.

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