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Two-dimensional crystallization of a membrane protein on a detergent-resistant lipid monolayer1
doi:10.1006/jmbi.2001.4629 available online at http://www.idealibrary.com on
J. Mol. Biol. (2001) 308, 639±647
Two-dimensional Crystallization of a Membrane Protein on a Detergent-resistant Lipid Monolayer Luc Lebeau1*, Franck Lach1, Catherine VeÂnien-Bryan2, Anne Renault3 Jens Dietrich4, Thomas Jahn5, Michael G. Palmgren5 Werner KuÈhlbrandt4 and Charles Mioskowski1* 1
Laboratoire de SyntheÁse Bioorganique associe au CNRS Universite Louis Pasteur 67401 Illkirch, France 2
Institut de Biologie Structurale CEA-CNRS, 38027 Genoble Cedex 1, France 3
Laboratoire de SpectromeÂtrie Physique, Universite Joseph Fourier, 38402 Saint Martin d'HeÁres, France 4 Max-Planck-Institut fuÈr Biophysik, 60528 Frankfurt am Main, Germany
Two-dimensional crystals of a membrane protein, the proton ATPase from plant plasma membranes, have been obtained by a new strategy based on the use of functionalized, ¯uorinated lipids spread at the airwater interface. Monolayers of the ¯uorinated lipids are stable even in the presence of high concentrations of various detergents as was established by ellipsometry measurements. A nickel functionalized ¯uorinated lipid was spread into a monolayer at the air-water interface. The overexpressed His-tagged ATPase solubilized by detergents was added to the subphase. 2D crystals of the membrane protein, embedded in a lipid bilayer, formed as the detergent was removed by adsorption. Electron microscopy indicated that the 2D crystals were single layers with dimensions of 10 mm or more. Image processing yielded a projection map at Ê resolution, showing three well-separated domains of the membrane9A embedded proton ATPase. # 2001 Academic Press
5
The Royal Veterinary and Agricultural University, 1871 Frederiksberg C, Copenhagen Denmark *Corresponding authors
Keywords: lipid layer crystallization; 2D crystal; membrane protein; nickel-chelating lipid; ¯uorinated lipid
Introduction Genome sequences indicate that roughly 20-40 % of the proteins in any living cell are membrane proteins (Frishman & Mewes, 1997; Jones, 1998). To date, the three-dimensional structures of approximately 10,000 soluble proteins have been determined by X-ray crystallography. However, only the structures of roughly 20 membrane proteins have been solved. To a large extent this is due to the dif®culty of growing well-ordered memPresent addresses: A. Renault, GMCM, BaÃt. 11A, Universite Rennes 1, Campus de Beaulieu, 35042 Rennes Cedex, France; C. VeÂnien-Bryan, Laboratory of Molecular Biophysics, South Parks Road, Oxford OX1 3QU, UK. Abbreviations used: His-tag, histadine tag; Ni-NTA, Ni2-chelating nitrilotriacetate; LE, liquid expanded. E-mail addresses of the corresponding authors: [email protected]; [email protected] 0022-2836/01/040639±9 $35.00/0
brane protein crystals in view of the amphiphilic nature of these macromolecules. The use of detergents (Helenius & Simons, 1975; Tanford & Reynolds, 1976) to mask the hydrophobic surface areas, and more recently the use of lipidic cubic phases (Landau & Rosenbusch, 1996) have yielded some excellent 3D crystals of membrane proteins. However despite these breakthroughs, progress in growing well-ordered 3D crystals of membrane proteins suitable for X-ray crystallographic analysis has been slow. As most membrane proteins are oligomeric complexes with a molecular weight greater than 50 kDa, even without accounting for bound detergent, they are well outside the reach of current solution NMR techniques. Alternative approaches to structure determination are essential, such as electron microscopy. This method relies on the ability to incorporate the membrane protein into a bilayer. Favourable protein-protein interactions may lead to large, well ordered 2D crystals suitable for structural studies by electron microscopy (Henderson et al., 1990; # 2001 Academic Press
640 KuÈhlbrandt et al., 1994). Even though the electron crystallographic method has proved successful for the determination of the molecular structure of many membrane proteins(Walz & Grigorieff, 1998) new more reliable crystallization methods are needed. In the case of soluble proteins, a high local protein concentration obtained by attaching the protein to monolayers of conventional, charged or functionalized lipids spread at an air-water interface (Jap et al., 1992; Lebeau et al., 1996). For example, soluble proteins carrying a histidine tag (His-tag) have been crystallized on monolayers of regular lipids derivatized with a Ni2-chelating nitrilotriacetate (Ni-NTA) group (Dietrich et al., 1995; Kubalek et al., 1994; VeÂnien-Bryan et al., 1997). Anchoring the protein onto the ®lm imposes a particular orientation of the macromolecule relative to the interface. This preferential orientation may be an important precondition for the subsequent formation of a 2D protein lattice. Efforts to extend the approach of 2D crystallization on lipid monolayers to membrane proteins were complicated by the characteristic tendency of detergents to solubilize monolayers of regular lipids (Figure 1A). In a recent study, 2D crystals of two histidine-tagged membrane proteins have been obtained on a monolayer of conventional lipids derivatized with a Ni2-chelating group (LeÂvy et al., 1999). Successful crystallization required precise conditions that included the use of low CMC detergent and a very high binding af®nity of the protein for the functionalized lipid in order to stabilize the interface (Lebeau et al., 1996). These two conditions seem to represent a limitation of this technique, and therefore this approach may not be of general use. We tackled the problem of solubilization of the lipid monolayer by designing partially ¯uorinated lipids which, when spread at the interface, display a high resistance toward solubilization by detergents. Fluorinated amphiphilic compounds, spread at an air-water interface form monolayers that are unperturbed by small hydrophobic compounds such as detergents (Held et al., 1997). This property of the ¯uorinated amphiphiles originates from the non-ideality of alkane/per¯uoroalkane mixtures (Mukerjee, 1994; Mukerjee & Yang, 1976). Indeed mixing water with ¯uorocarbon and hydrocarbon compounds yields a three-phase-system as none of the components are miscible. We anticipated that this phenomenon could be exploited to prepare lipid layers that might not be solubilized by conventional detergents and hence would be useful for 2D membrane protein crystallization (Figure 1(b)). A nickel-chelating moiety was introduced into the polar head group of the ¯uorinated lipid (F. Lach et al., unpublished results), thus providing a speci®c site for strong interaction with a polyhistidine tag on the membrane protein. Most cells contain only small amounts of a particular membrane protein. Production of large quantities of a His-tagged membrane protein is
Membrane Protein 2D Crystallization on Lipid Layers
Figure 1. (a) Regular non-¯uorinated lipid layers are not stable upon addition of detergents and are solubilized rapidly as mixed micelles. (b) When speci®cally designed ¯uorinated lipids are spread at the air-water interface, the lipid layer remains stable for hours in the presence of detergents. This allows a membrane protein to bind and orient at the interface. Subsequent removal of detergent by adsorption leads to the reconstitution of the protein in a membrane and the possible formation of protein 2D arrays.
Membrane Protein 2D Crystallization on Lipid Layers
achieved by overexpression. The puri®cation step then takes place using Ni2-chelating chromatography as immobilized Ni2 bind tightly to the poly-histidine tags carried by the expressed protein (Hochuli et al., 1987). Here, we show that His-tags can also be employed for the 2D crystallization of membrane proteins. Using this new approach we obtained very large, well-ordered 2D crystals of a plasma membrane H-ATPase. Image processing of electron cryo-micrographs yielded a projection Ê resolution. map at 9 A
Results and Discussion Resistance of fluorinated lipids to the solubilization by detergents The structures of the synthetic lipids used in these experiments are shown in Figure 2. Details of the synthesis will be reported elsewhere. Lipids 1 and 2 differ by the length of the linker between the hydrophobic section and the NTA functional group. A longer linker was thought to give more freedom of movement and thus to be conducive to a better crystal packing of the protein Lipid 3 was used as a diluting lipid matrix. Lipid 4 is the regular non-¯uorinated lipid used for comparative studies. Monolayer properties of the lipids 1 and 2 were investigated by mechanical and optical techniques. Isothermal surface pressure versus area curves
Figure 2. Structure of the partially ¯uorinated Ni2chelating lipids used in this study. The lipids were designed to spread at the air-water interface into monolayers in the ¯uid LE (liquid expanded) phase. Lipids 1 and 2 differ by the length of the linker between the fatty chains and the NTA moiety. Lipid 3 was used as a diluting lipid in the crystallization experiments. Lipid 4 (Ni-NTA-DOGA) was used for comparative experiments.
641 were determined using a Langmuir trough equipped with an electrobalance. Pro®les indicate that the two ¯uorinated lipids form stable monolayers at the air-water interface (Figure 3). It is well known the lipid matrix must be spread in the ¯uid liquid-expanded (LE) phase for successful 2D crystallization of proteins on lipid monolayers (Lebeau et al., 1996). This is essential to allow in-plane motions of the lipid-protein complexes so they can diffuse laterally and possibly organize into 2D arrays. An adequate ¯uidity of the lipid layers is ensured by branched aliphatic chains, which prevent tight, liquid-crystalline packing (Menger et al., 1988). On the pressure versus area curves, a plateau is observed at 1-2 nm2/molecule that is interpreted as the result of a conformational change of the branched aliphatic chains during monolayer compression in the ¯uid phase. The stability of the ¯uorinated monolayers in the presence of detergents was monitored by ellipsometry, an optical technique which is especially suited to the study of liquid surfaces. It is a non-invasive method that is sensitive to the density and thickness of the interface layer (Berge & Renault, 1993). The technique has been used to investigate the adsorption of proteins onto lipid monolayers (Andree et al., 1990; Caide et al., 1996; VeÂnien-Bryan et al., 1998). The ellipsometric angle d correlates with the amount of lipid and detergent adsorbed at the interface and is taken to be zero for a pure air-water interface. Spreading the non-¯uorinated nickel-chelating lipid 4 (Ni-NTADOGA, Figure 2) at the interface to form a monolayer produces a shift in d to 21 (Figure 4(a), arrow 1). The monolayer proves to be perfectly stable. If detergent is added in the aqueous phase (Figure 4(a), arrow 2), the value for the ellipsometric angle decreases within seconds and the ellipsometric angle reaches d 2(0.5) . This ellipsometric angle represents the value obtained for a
Figure 3. Isothermal surface pressure vs. lipid molecular area curves obtained with lipids 1 (thin line) and 2 (bold line) spread at the air-water interface at 20 C. Experimental procedures are described elsewhere (Lebeau et al., 1988).
642
Figure 4. Effect of detergents on the lipid monolayers monitored by ellipsometry. (a) Monolayer made made of non-¯uorinated lipid 4 (Ni-NTA-DOGA): the lipid is spread into a monolayer at the interface (arrow 1), then detergent (Emulgen 911, 0.5 CMC) is added into the aqueous subphase (arrow 2). (b)-(g) Monolayers made of ¯uorinated lipid 2. The experiment in (b) is similar to that described in (a): the lipid is spread at the interface (arrow 1), then Emulgen 911 is sequentially added into the aqueous subphase (arrow 2: 0.5 CMC; arrow 3: 0.5 CMC; arrow 4: 0.5 CMC; arrow 5: 1.5 CMC; arrow 6: 2.0 CMC). Various detergents have been tested: (c) Emulgen 911; (d) dodecyl-b-D-maltoside; (e) CHAPS; (f) SDS; (g) octyl-b-D-glucopyranoside. Arrows indicate addition of detergent (0.5 CMC). The loose dashed line refers to the value of the ellipsometric angle d recorded after addition of the corresponding detergent with no ¯uorinated lipid spread at the interface. The close dashed line indicates the equilibrium value for d when no detergent is added.
Membrane Protein 2D Crystallization on Lipid Layers
turbation at the interface. After a few minutes however the system goes back to equilibrium and d stabilizes close to the initial value of 7 . Repeated additions of detergent (Figure 4(b): arrow 3: 0.5 CMC; arrow 4: 0.5 CMC; arrow 5: 1.5 CMC; arrow 6: 2.0 CMC) to a ®nal ®vefold CMC detergent concentration only transiently disturbs the monolayer. Therefore the detergent Emulgen 911 is unsuccessful at solubilizing the monolayer of lipid 2. The ellipsometric angle remains constant for detergent Emulgen 911 concentration up to 5 CMC over a period of days. The stability of the ¯uorinated monolayer was investigated with detergents that have an extended range of CMC values (Figure 4(c)-(g)). High CMC detergents (octyl-b-D-glucopyranoside, CMC 7.3 g/l; sodium dodecyl sulfate, CMC 2.3 g/l) do not destabilize the ¯uorinated matrix more than those with lower CMC values (Emulgen 911, CMC 0.2 g/l; dodecyl-b-Dmaltoside, CMC 0.08 g/l). Some minor deviations of the d value can be interpreted as partial accumulation of detergent molecules at the polar interface created by the head groups of the nickelchelating lipids. This effect is especially noticeable with the CHAPS detergent (Figure 4(e)). It is possible for the sulphonate moiety in this detergent molecule to displace one or both water molecules from the nickel head group of lipid 2. This would then allow one or two detergent molecules to be tethered at the interface by one nickel ion, modifying the ellipsometric signal. But in no case are the ¯uorinated lipids removed from the interface and solubilized by the detergents. Lipid 1 exhibits very similar properties (data not shown). In some cases we have noticed that the ellipsometric signal obtained with the ¯uorinated lipids is more noisy than the signal obtained with the non-¯uorinated lipids. This is particularly visible in Figure 4(b). These ¯uctuations are interpreted as the presence of some heterogeneous lipid domains moving at the air-water interface. It is interesting to note that these ¯uctuations disappear when detergent is added (Figure 4). We conclude from these experiments that the partially ¯uorinated lipids spread at the air-water interface are resistant to solubilization by a large variety of detergents. Formation of 2D crystals using the fluorinated lipid monolayers
water-detergent solution. This indicates that the addition of even a small quantity of detergent (0.5 CMC) to the subphase under the lipid monolayer results in the complete and rapid solubilization of the lipid at the interface. The same experiment with the ¯uorinated lipid 1 or 2 takes a totally different course. Deposition of the ¯uorinated lipid at the air-water interface increases the d value to 7(1) (Figure 4(b), arrow 1). The addition of Emulgen 911 (Figure 4(b) arrow 2: 0.5 CMC) in the aqueous subphase provokes a transient per-
In order to check that the newly designed lipids were good candidates for promoting the formation of protein 2D crystals, we ®rst tested the lipids with streptavidin. This soluble protein has been extensively studied and the crystallization conditions are well known (Blankenburg et al., 1989). In this experiment, the partially ¯uorinated lipid was functionalized with a biotinyl group which binds tightly to streptavidin, rather than with the Ni-NTA group. Crystals of streptavidin were obtained reproducibly (data not shown), providing evidence that these ¯uorinated lipids possess
Membrane Protein 2D Crystallization on Lipid Layers
the correct ¯uidity and mobility properties at the interface necessary for the formation of crystals. We then used the ¯uorinated lipids functionalized with Ni-NTA moiety in order to crystallize membrane proteins. The plasma membrane H-ATPase (AHA2) from Arabidopsis thaliana is a single-subunit integral membrane protein with a molecular mass of 104 kDa. It belongs to the family of P-type transport ATPases. These are widely distributed biological energy transducers that convert the free energy resulting from ATP hydrolysis into an electrochemical ion gradient across the membrane (KuÈhlbrandt et al., 1998). Two-dimensional crystals of the corresponding enzyme from the bread mold Neurospora crassa have been obtained recently by surface crystallization on a solid carbon ®lm support. Well-ordered 2D crystals of the dodecylb-D-maltoside protein complex were found to grow rapidly in the presence of polyethylene glycol by a procedure based on that developed for growing 3D crystals (Cyrklaff et al., 1995). Even though the protein was not embedded in a lipid bilayer, the 2D crystals proved useful for structure analysis to Ê resolution (Auer et al., 1998, 1999). Two-dimen8A sional of the Neurospora crassa H-ATPase by conventional reconstitution into lipid membranes has so far been unsuccessful. The plant plasma membrane ATPase from A. thaliana was expressed in yeast with a C-terminal His-tag and puri®ed by Ni2-chelating chromatography. With the ¯uorinated lipids spread at the air-water interface, large well-ordered 2D crystals of the H-ATPase formed reproducibly. Crystals were obtained by spreading a mixture of lipids 2 and 3 at a ratio of 1:6 (w/w) onto 50 ml of the crystallization solution containing the protein and membrane lipids, both solubilized in dodecylb-D-maltoside. The drop was left at room temperature for a period of 24 hours during which the protein adsorbed to the ¯uorinated lipid layer. After this time, detergent was removed by adding 8-10 Bio-Beads1, followed by incubation for another 24-hour period to enable the crystal lattice to form. The Ni-NTA head group caused the protein to concentrate and orient at the interface between ¯uorinated lipid and water even when using a dilute protein solution (0.15 mg/ml). When samples were transfered to electron microscopy grids before the detergent removal step, large noncrystalline sheets of protein were observed (data not shown). At this stage the protein has attached to the lipid layer, probably as tightly clustered single molecules or oligomers while still being surrounded by detergent. Thus the crystal lattice clearly does not form before removal of detergent and protein insertion into a lipid bilayer. Complementary control experiments conducted in the presence of 250 mM imidazole or 150 mM NiSO4 under otherwise identical conditions did not produce crystals or even sheets of disordered protein (data not shown). As expected, both
643 chemicals were effective inhibitors of protein binding. Imidazole is used routinely in Ni2-chelating chromatography release to bound protein from the columns. Low concentrations of NiSO4 were suf®cient to saturate the Ni2-chelating histidine tags on the protein so that it was no longer able to attach to the derivatized lipid. These observations provide conclusive evidences that attachment of the protein via its His-tag to the Ni-NTA-derivatized lipid was an essential prerequisite for the formation of large, single layer crystals. For electron microscopy the 2D crystals were picked up from the surface of the drop with a carbon coated copper grid. Crystals were either negatively stained or prepared for electron cryomicroscopy by blotting, plunging into liquid ethane and mounting in a cryo-transfer holder. Grids were examined in an electron microscope and micrographs were recorded for structure determination by image analysis. Large membrane sheets, typically measuring more than 10 mm across, were frequently found and consisted of either a single crystal lattice, or of a few lattices fused in plane, as revealed by electron microscopy of negatively stained crystals (Figure 5). Projection map and structure of the H-ATPase Electron micrographs of frozen-hydrated 2D crystals were selected for coherent, well-ordered
Figure 5. 2D crystals of the Arabidopsis thaliana plasma membrane H-ATPase grown on a layer of lipids 2 and 3 after removal of the detergent, dodecyl-b-D-maltoside, by adsorption to Bio-Beads1. Crystals were stained with 2 % (w/w) uranyl acetate as a contrasting agent. (a) Overview of a large crystalline area. (b) The rectangular crystal lattice is visible at higher magni®cation.
644
Membrane Protein 2D Crystallization on Lipid Layers
lattices by optical diffraction, digitized and further analyzed by image processing (Crowther et al., 1996; Henderson et al., 1986). The unit cell parÊ , b 139 A Ê ameters were calculated as: a 151 A and g 90 (Table 1). Phase comparisons indicated that the crystal symmetry was P22121. Upon correction of lattice distortions, amplitudes and phases Ê resolution (Figure 6(a)). The were obtained to 9 A projection map (Figure 6(b)) looks different from the corresponding map of the Neurospora enzyme Ê . While the latter at a comparable resolution of 8 A crystallizes as a hexamer, the plant ATPase apparently crystallizes as a dimer. Each monomer consists of three distinct areas of density of roughly equal size in projection whereas the Neurospora ATPase does not show such a clear division into three density regions. The two enzymes are closely related by homology of the polypeptide sequence except that the N and C termini have different lengths (Axelsen & Palmgren, 1998). The projection maps must re¯ect these structural differences, but in addition may represent different conformations of the enzyme. This would not be surprising as the comparison of the Neurospora H-ATPase with the related Ca2-ATPase from sarcoplasmic reticulum has shown a high degree of conformational ¯exibility in the large cytoplasmatic domains, apparently associated with the E1 to E2 transition in the ion pumping cycle (Stokes et al., 1999). Such conformational changes could easily account for the observed differences in the projection maps. The P22121 symmetry of the 2D crystals of the plant H-ATPase indicates that the up and down orientation of each dimer alternates with respect to the plane of the lipid monolayer. This suggests that only one half of the molecules make speci®c contacts through their His-tag with the Ni-NTA moiety of the lipid head groups. The other half packs into the crystal in the upside-down orientation, resulting in a head-to-tail arrangement of pairs of ATPase molecules related by the 2-fold screw axes in the plane of the membrane. Small vesicular 2D crystals of the plant proton ATPase expressed in Saccharomyces cerevisiae have
been obtained by conventional detergent dialysis in the presence of membrane lipids (T.J., M.G.P., J.D., W.K. et al., unpublished results). It is interesting that these crystals have the same unit cell dimensions and symmetry as the crystalline sheets forming on lipid monolayers described here. They remain however much smaller, with diameters rarely exceeding 1-2 mm. Electron micrographs of the ATPase vesicle crystals always show two superimposed crystal lattices, one from each layer, whereas the 2D crystals described herein are single sheets, with only one lattice visible in the electron microscope. The fact that both crystal types show the same packing implies that the ATPase is embedded in a lipid bilayer both in the vesicle crystals and in the large crystalline sheets reported here. We cannot rule out that the crystalline sheets are produced by fusion of pre-formed crystalline vesicles which attach via their His-tags to the monolayers of ¯uorinated lipids. We think this mechanism of crystal formation is unlikely however because the protein clearly attaches to the monolayer prior to lattice formation, resulting in large, unordered layers which are not observed under conditions which prevent protein attachment. General use of fluorinated lipids for the 2D crystallization of membrane proteins Although it may be possible to grow 2D crystals either by conventional lipid reconstitution or on monolayers of detergent-resistant lipids, our new approach clearly has a number of important advantages for structure determination of membrane proteins by electron crystallography. The technique is quick and reproducible, and requires only minimal amounts of protein and functionalized lipid (approximately 7.5 mg and 500 nmol respectively per trial). The 2D crystals we obtained are large, single layers, offering the potential of electron diffraction as a straightforward test of crystalline order and state of preservation. This will make it much easier to optimize crystallization
Table 1. Electron crystallographic data Plane group symmetry Unit cell dimensions Number of images and lattices Range of defocus Total number of observations Number of unique observations Ê Overall phase residual to 9 A Ê )a Resolution range (A >20.4 20.4-14.3 14.3-11.6 11.6-10.1 10.1-9.0
P22121 Ê ; b138.9(1.5) A Ê ; g90 a150.9(1) A 5 Ê 7400-9500 A 611 191 25.5 Number of unique reflections Phase residualb 39 13.2 40 23.4 38 29.8 36 30.3 38 31.4
a Phase residual of re¯ections in resolution ranges, showing the mean deviation of averaged phases from the theoretical value, 0 or 180 . b Random data would give a value of 45 .
645
Membrane Protein 2D Crystallization on Lipid Layers
conditions for obtaining large, highly ordered 2D crystals and to collect data by electron diffraction and image processing. Finally, the approach described here may be applied to any membrane protein that can be expressed with a His-tag, and thus offers the prospect of a much needed rational, conceptually simple and potentially general method for the 2D crystallization of membrane proteins.
Materials and Methods Lipids, detergents and protein Phospholipids of the highest purity available were purchased from Avanti Polar-Lipids. Dodecyl-b-D-maltoside was from Calbiochem and Bio-Beads1 SM2 (20-50 mesh) from Bio-Rad. Ni-NTA-DOGA was prepared as described by (VeÂnien-Bryan et al. (1997). The synthesis of lipids 1, 2, and 3 will be described in detail elsewhere. Lipid stock solutions were kept at 1 or 2 mg/ml in chloroform/methanol (1:1, v/v) at ÿ20 C under argon gas. His-tagged H-ATPase was over-expressed and puri®ed to homogeneity as described (Jahn et al., 2001). Ellipsometry
Figure 6. (a) Structure factors of a 2D crystal of the Arabidopsis plasma membrane H-ATPase. The image was obtained by electron cryo-microscopy and image processing of unstained 2D crystals. The size of the square and the number re¯ect the signal-to-noise ratio of the computed Fourier component (Henderson et al., 1986). The concentric rings indicate the zero positions of the contrast transfer function. The resolution at the edge Ê . (b) Projection map of the plasma of the plot is 9 A Ê resolution calculated from membrane H-ATPase at 9 A merged amplitudes and phases from ®ve independent crystals with P22121 symmetry. Solid lines indicate density above the mean. Negative contours are indicated by dotted lines. The unit cell contains four molecules and Ê , b 139 A Ê and g 90 . The parameters are a 151 A monomer boundary cannot be de®ned with certainty in the projection map but comparison with the projected structure of the Neurospora enzyme suggests the outlined shape of the monomer.
The ellipsometric measurements were carried out with a conventional ellipsometer using a He-Ne laser operating at 632.8 nm as described by Berge & Renault (1993) and VeÂnien-Bryan et al. (1998). After re¯ection at the surface of water, the beam is analysed. The phase difference between the vertical and horizontal beam polarizations and so the ellipsometric angle (d) are determined. The variation of the ellisometric angle is a relevant probe for changes occurring at the interface. It is proportional to the amount of lipid and detergent adsorbed at the interface. The sample cell is made of Te¯on1 and has a volume of 8 ml. Basically, a lipid monolayer was spread at the air-water interface. Once equilibrium was reached, detergent was injected into the subphase. Homogeneity of the solution was ensured using a peristaltic pump. When the surface tension was measured in the same trough, we used the Wilhelmy method using a 1-cmwide ®lter paper. For more precise measurements of the surface pressure versus lipid molecular area we used a Wilhelmy plate coupled to a linear transducer (KSV 9900 Surface Barostat, KSV, Finland) and a Langmuir Te¯on1 trough (165 mm 270 mm). Monolayers were spread from 0.5 mM lipid solutions. Volume ranging from 100 to 300 ml were delivered at several locations across the water surface with a Hamilton microsyringe. At least 15 minutes were allowed for the spreading solvent to evaporate before monolayer compression. The subphase consisted of puri®ed water (pH 5.35, Millipore1 ®ltration system, Barnstead, NANO pure 11) with a resistivity of 18.2 106 cm. Surface pressure versus molecular area curves were recorded at 22.0(0.2) C with a comÊ 2 per lipid molecule per minute. pression speed of 2-5 A All measurements were carried out in triplicate. Protein binding and crystallization The protein (150 mg/ml) and egg phosphatidylcholine/dodecyl-b-D-maltoside mixed micelles (lipid to protein ratio 1:3, w/w) were placed in a Te¯on1 well (6 mm diameter, 1 mm deep) in a buffer containing 20 % (w/w) glycerol, 20 mM Mes, 150 mM KCl and a ®nal
646 detergent concentration of about 0.1 % (w/w) dodecylb-D-maltoside for membrane reconstitution. The resulting drop (50 ml) was coated with a monolayer of ¯uorinated lipid by adding 1 ml of the working solution (lipids 2 and 3 at a ratio of 1:6, w/w, 500 mM in chloroform/hexane 1:1, v/v) to the surface with a Hamilton syringe. After a ®rst 24-hour incubation period at 20 C in a humid chamber, eight to ten Bio-Beads1 were added to the well through a submerged channel, to avoid any disturbance of the drop surface. Two-dimensional crystals were allowed to form for another 24 hours and were transferred onto electron microscopy grids for analysis. Addition of 1 mM EDTA and 0.25 mM DTT to the subphase to stabilize the protein did not interfere with the formation of crystalline monolayers. These conditions were similar to those used routinely for column chromatography of the His-tagged ATPase on Ni2-chelating columns (M.J., M.G.P., J.D., W.K. et al., unpublished results). Control experiments in the presence of 10 mM EDTA, 150 mM NiSO4 or 250 mM imidazole which would release the His-tagged protein from the NTA column did not yield any protein crystalline sheets. Electron microscopy and image analysis Carbon-coated electron microscope grids were baked at 150 C for one hour prior to use in order to enhance the adhesion of the crystals by making the grids more hydrophobic. The crystals were picked up by placing a carbon-coated grid on top of the crystallization well for several minutes. The grid was removed, blotted with ®lter paper and either negatively stained with 2 % (w/w) uranyl acetate for 30 seconds or, plunged into liquid ethane and transferred to a GATAN cryo-transfer stage. Images were recorded on a Philips CM 120 transmission electron microscope operating at 120 kV in the low dose mode at a magni®cation of 45,000 . Selected images were processed using the MRC program suite (Crowther et al., 1996; Henderson et al., 1986).
Acknowledgements We are grateful to Declan Doyle for his critical reading of the manuscript and constructive comments. We thank the AFM (France) for a grant to F.L. The EU Biotechnology Program is acknowledged for ®nancial support (M.G.P. and W.K.).
References Andree, H. A. M., Reutelingsperger, C. P. M., Hauptmann, R., Hemker, H. C., Hermens, W. T. & Willems, G. M. (1990). Binding of vascular anticoagulant a (VAC a) to planar phospholipid bilayers. J. Biol. Chem. 265, 4923-4928. Auer, M., Scarborough, G. A. & KuÈhlbrandt, W. (1998). Three-dimensional map of the plasma membrane H-ATPase in the open conformation. Nature, 392, 840-841. Auer, M., Scarborough, G. A. & KuÈhlbrandt, W. (1999). Surface crystallization of the plasma membrane H-ATPase on a carbon support ®lm for electron crystallography. J. Mol. Biol. 287, 961-968. Axelsen, K. B. & Palmgren, M. G. (1998). Evolution of substrate speci®cities in the P-type ATPase superfamily. J. Mol. Evol. 46, 84-101.
Membrane Protein 2D Crystallization on Lipid Layers Berge, B. & Renault, A. (1993). Ellipsometry study of 2D crystallization of 1-alcohol monolayers at the water interface. Europhys. Letters, 21, 773-777. Blankenburg, R., Meller, P., Ringsdorf, H. & Salesse, C. (1989). Interaction between biotin lipids and streptavidin in monolayers: formation of oriented twodimentionsal protein domains induced by surface recognition. Biochemistry, 28, 8214-8221. Caide, X., Liu, Z., Gao, Q. F., Zhou, Q. Z. & Sui, S. F. (1996). Speci®c interaction of rabbit C-reactive protein with phospholipid membranes. Thin Solid Films, 285, 793-796. Crowther, R. A., Henderson, R. & Smith, J. M. (1996). MRC Image processing programs. J. Struct. Biol. 116, 9-16. Cyrklaff, M., Auer, M., KuÈhlbrandt, W. & Scarborough, G. A. (1995). 2-D structure of Neurospora crassa plasma membrane ATPase as determined by electron cryomicroscopy. EMBO J. 14, 1854-1857. Dietrich, C., Schmitt, L. & TampeÂ, R. (1995). Molecular organization of histidine-tagged biomolecules at self-assembled lipid interfaces using a novel class of chelator lipids. Proc. Natl Acad. Sci. USA, 92, 90149018. Frishman, D. & Mewes, H. W. (1997). Protein structural classes in ®ve complete genomes. Nature Struct. Biol. 4, 626-628. Held, P., Lach, F., Lebeau, L. & Mioskowski, C. (1997). Synthesis and preliminary evaluation of a new class of ¯uorinated amphiphiles designed for in-plane immobilization of biological macromolecules. Tetrahedron Letters, 38, 1937-1940. Helenius, A. & Simons, K. (1975). Solubilization of membrane proteins by detergents. Biochim. Biophys. Acta, 415, 29-79. Henderson, R., Baldwin, J. M., Downing, K. H., Lepault, J. & Zemlin, F. (1986). Structure of purple membrane from Halobacterium halobium: recording, measurement and evaluation of electron microÊ resolution. Ultramicroscopy, 19, 147graphs at 3.5 A 178. Henderson, R., Baldwin, J. M., Ceska, T. A., Zemlin, F., Beckmann, E. & Downing, K. H. (1990). Model for the structure of bacteriorhodopsin based on highresolution electron cryomicroscopy. J. Mol. Biol. 213, 899-929. Hochuli, E., DoÈbeli, H. & Schacher, A. (1987). New metal chelate adsorbent selective for proteins and peptides containing neighbouring histidine residues. J. Chromatog. 411, 177-184. Jahn, T., Dietrich, J., Andersen, B., Leidvik, B., Otter, C., Briving, C., KuÈhlbrandt, W. & Palmgren, M. G. (2001). Large scale expression, puri®cation and 2D crystallization of recombinant plant plasma membrane HATPase. J. Mol. Biol. In the press. Jap, B. K., Zulauf, M., Scheybani, T., Hefti, A., Baumeister, W., Aebi, U. & Engel, A. (1992). 2D crystallization: from art to science. Ultramicroscopy, 46, 45-84. Jones, D. T. (1998). Do transmembrane protein superfolds exist? FEBS Letters, 423, 281-285. Kubalek, E. W., Le Grice, S. F. J. & Brown, P. O. (1994). Two-dimensional crystallization of histidine-tagged HIV-1 reverse transcriptase promoted by a novel nickel-chelating lipid. J. Struct. Biol. 113, 117-123. KuÈhlbrandt, W., Wang, D. N. & Fujiyoshi, Y. (1994). Atomic model of plant light-harvesting complex by electron crystallography. Nature, 367, 614-621.
Membrane Protein 2D Crystallization on Lipid Layers KuÈhlbrandt, W., Auer, M. & Scarborough, G. A. (1998). Structure of the P-type ATPases. Curr. Opin. Struct. Biol. 8, 510-516. Landau, E. M. & Rosenbusch, J. P. (1996). Lipidic cubic phases: a novel concept for the crystallization of membrane proteins. Proc. Natl Acad. Sci. USA, 93, 14532-14535. Lebeau, L., Mioskowski, C. & Oudet, P. (1988). Systematic study of phospholipids linked to a steroid derivative, spread into a monolayer at the air/ water interface. Biochim. Biophys. Acta, 939, 417-429. Lebeau, L., Schultz, P., CeÂlia, H., MeÂsini, P., Nuss, S. & Klinger, C. et al. (1996). Speci®cally designed lipid assemblies as tools for two-dimensional crystallization of soluble biological macromolecules. In Handbook of Non-medical Applications of Liposomes (Barenholz, Y. & Lasic, D. D., eds), vol. II pp. 153186, CRC Press, Boca Raton. LeÂvy, D., Mosser, G., Lambert, O., Moeck, G. S., Bald, D. & Rigaud, J. L. (1999). Two-dimensional crystallization on lipid layer: a successful approach for membrane proteins. J. Struct. Biol. 127, 44-52. Menger, F. M., Wood, M. G. J., Richardson, S., Zhou, Q., Elrington, A. R. & Sherrod, M. J. (1988). Chainsubstituted lipids in monolayer ®lms. A study of molecular packing. J. Am. Chem. Soc. 110, 6797-6803. Mukerjee, P. (1994). Fluorocarbon-hydrocarbon interactions in micelles and other lipid assemblies, at interfaces, and in solutions. Colloids Surf. A, 84, 1-10.
647 Mukerjee, P. & Yang, A. Y. S. (1976). Non-ideality of mixing of micelles of ¯uorocarbon and hydrocarbon surfactants and evidence of partial miscibility from differential conductance data. J. Phys. Chem. 80, 1388-1390. Stokes, D. L., Auer, M., Zhang, P. & KuÈhlbrandt, W. (1999). Comparison of H-ATPase and Ca2-ATPase suggests that a large conformational change initiates P-type ion pump reaction cycles. Curr. Biol. 9, 672679. Tanford, C. & Reynolds, J. A. (1976). Characterization of membrane proteins in detergent solutions. Biochim. Biophys. Acta, 457, 133-170. VeÂnien-Bryan, C., Balavoine, F., Toussaint, B., Mioskowski, C., Hewat, E. A., Helme, B. & Vignais, P. M. (1997). Structural study of the response regulator HupR from Rhodobacter capsulatus. Electron microscopy of two-dimensional crystals on a nickelchelating lipid. J. Mol. Biol. 274, 687-692. VeÂnien-Bryan, C., Lenne, P. F., Zakri, C., Renault, A., Brisson, A., Legrand, J. F. & Berge, B. (1998). Characterization of the growth of 2D protein crystals on a lipid monolayer by ellipsometry and rigidity measurements coupled to electron microscopy. Biophys. J. 74, 2649-2657. Walz, T. & Grigorieff, N. (1998). Electron crystallography of two-dimensional crystals of membrane proteins. J. Struct. Biol. 121, 142-161.
Edited by R. Huber (Received 8 November 2000; received in revised form 16 February 2001; accepted 12 March 2001)
J. Mol. Biol. (2001) 308, 639±647
Two-dimensional Crystallization of a Membrane Protein on a Detergent-resistant Lipid Monolayer Luc Lebeau1*, Franck Lach1, Catherine VeÂnien-Bryan2, Anne Renault3 Jens Dietrich4, Thomas Jahn5, Michael G. Palmgren5 Werner KuÈhlbrandt4 and Charles Mioskowski1* 1
Laboratoire de SyntheÁse Bioorganique associe au CNRS Universite Louis Pasteur 67401 Illkirch, France 2
Institut de Biologie Structurale CEA-CNRS, 38027 Genoble Cedex 1, France 3
Laboratoire de SpectromeÂtrie Physique, Universite Joseph Fourier, 38402 Saint Martin d'HeÁres, France 4 Max-Planck-Institut fuÈr Biophysik, 60528 Frankfurt am Main, Germany
Two-dimensional crystals of a membrane protein, the proton ATPase from plant plasma membranes, have been obtained by a new strategy based on the use of functionalized, ¯uorinated lipids spread at the airwater interface. Monolayers of the ¯uorinated lipids are stable even in the presence of high concentrations of various detergents as was established by ellipsometry measurements. A nickel functionalized ¯uorinated lipid was spread into a monolayer at the air-water interface. The overexpressed His-tagged ATPase solubilized by detergents was added to the subphase. 2D crystals of the membrane protein, embedded in a lipid bilayer, formed as the detergent was removed by adsorption. Electron microscopy indicated that the 2D crystals were single layers with dimensions of 10 mm or more. Image processing yielded a projection map at Ê resolution, showing three well-separated domains of the membrane9A embedded proton ATPase. # 2001 Academic Press
5
The Royal Veterinary and Agricultural University, 1871 Frederiksberg C, Copenhagen Denmark *Corresponding authors
Keywords: lipid layer crystallization; 2D crystal; membrane protein; nickel-chelating lipid; ¯uorinated lipid
Introduction Genome sequences indicate that roughly 20-40 % of the proteins in any living cell are membrane proteins (Frishman & Mewes, 1997; Jones, 1998). To date, the three-dimensional structures of approximately 10,000 soluble proteins have been determined by X-ray crystallography. However, only the structures of roughly 20 membrane proteins have been solved. To a large extent this is due to the dif®culty of growing well-ordered memPresent addresses: A. Renault, GMCM, BaÃt. 11A, Universite Rennes 1, Campus de Beaulieu, 35042 Rennes Cedex, France; C. VeÂnien-Bryan, Laboratory of Molecular Biophysics, South Parks Road, Oxford OX1 3QU, UK. Abbreviations used: His-tag, histadine tag; Ni-NTA, Ni2-chelating nitrilotriacetate; LE, liquid expanded. E-mail addresses of the corresponding authors: [email protected]; [email protected] 0022-2836/01/040639±9 $35.00/0
brane protein crystals in view of the amphiphilic nature of these macromolecules. The use of detergents (Helenius & Simons, 1975; Tanford & Reynolds, 1976) to mask the hydrophobic surface areas, and more recently the use of lipidic cubic phases (Landau & Rosenbusch, 1996) have yielded some excellent 3D crystals of membrane proteins. However despite these breakthroughs, progress in growing well-ordered 3D crystals of membrane proteins suitable for X-ray crystallographic analysis has been slow. As most membrane proteins are oligomeric complexes with a molecular weight greater than 50 kDa, even without accounting for bound detergent, they are well outside the reach of current solution NMR techniques. Alternative approaches to structure determination are essential, such as electron microscopy. This method relies on the ability to incorporate the membrane protein into a bilayer. Favourable protein-protein interactions may lead to large, well ordered 2D crystals suitable for structural studies by electron microscopy (Henderson et al., 1990; # 2001 Academic Press
640 KuÈhlbrandt et al., 1994). Even though the electron crystallographic method has proved successful for the determination of the molecular structure of many membrane proteins(Walz & Grigorieff, 1998) new more reliable crystallization methods are needed. In the case of soluble proteins, a high local protein concentration obtained by attaching the protein to monolayers of conventional, charged or functionalized lipids spread at an air-water interface (Jap et al., 1992; Lebeau et al., 1996). For example, soluble proteins carrying a histidine tag (His-tag) have been crystallized on monolayers of regular lipids derivatized with a Ni2-chelating nitrilotriacetate (Ni-NTA) group (Dietrich et al., 1995; Kubalek et al., 1994; VeÂnien-Bryan et al., 1997). Anchoring the protein onto the ®lm imposes a particular orientation of the macromolecule relative to the interface. This preferential orientation may be an important precondition for the subsequent formation of a 2D protein lattice. Efforts to extend the approach of 2D crystallization on lipid monolayers to membrane proteins were complicated by the characteristic tendency of detergents to solubilize monolayers of regular lipids (Figure 1A). In a recent study, 2D crystals of two histidine-tagged membrane proteins have been obtained on a monolayer of conventional lipids derivatized with a Ni2-chelating group (LeÂvy et al., 1999). Successful crystallization required precise conditions that included the use of low CMC detergent and a very high binding af®nity of the protein for the functionalized lipid in order to stabilize the interface (Lebeau et al., 1996). These two conditions seem to represent a limitation of this technique, and therefore this approach may not be of general use. We tackled the problem of solubilization of the lipid monolayer by designing partially ¯uorinated lipids which, when spread at the interface, display a high resistance toward solubilization by detergents. Fluorinated amphiphilic compounds, spread at an air-water interface form monolayers that are unperturbed by small hydrophobic compounds such as detergents (Held et al., 1997). This property of the ¯uorinated amphiphiles originates from the non-ideality of alkane/per¯uoroalkane mixtures (Mukerjee, 1994; Mukerjee & Yang, 1976). Indeed mixing water with ¯uorocarbon and hydrocarbon compounds yields a three-phase-system as none of the components are miscible. We anticipated that this phenomenon could be exploited to prepare lipid layers that might not be solubilized by conventional detergents and hence would be useful for 2D membrane protein crystallization (Figure 1(b)). A nickel-chelating moiety was introduced into the polar head group of the ¯uorinated lipid (F. Lach et al., unpublished results), thus providing a speci®c site for strong interaction with a polyhistidine tag on the membrane protein. Most cells contain only small amounts of a particular membrane protein. Production of large quantities of a His-tagged membrane protein is
Membrane Protein 2D Crystallization on Lipid Layers
Figure 1. (a) Regular non-¯uorinated lipid layers are not stable upon addition of detergents and are solubilized rapidly as mixed micelles. (b) When speci®cally designed ¯uorinated lipids are spread at the air-water interface, the lipid layer remains stable for hours in the presence of detergents. This allows a membrane protein to bind and orient at the interface. Subsequent removal of detergent by adsorption leads to the reconstitution of the protein in a membrane and the possible formation of protein 2D arrays.
Membrane Protein 2D Crystallization on Lipid Layers
achieved by overexpression. The puri®cation step then takes place using Ni2-chelating chromatography as immobilized Ni2 bind tightly to the poly-histidine tags carried by the expressed protein (Hochuli et al., 1987). Here, we show that His-tags can also be employed for the 2D crystallization of membrane proteins. Using this new approach we obtained very large, well-ordered 2D crystals of a plasma membrane H-ATPase. Image processing of electron cryo-micrographs yielded a projection Ê resolution. map at 9 A
Results and Discussion Resistance of fluorinated lipids to the solubilization by detergents The structures of the synthetic lipids used in these experiments are shown in Figure 2. Details of the synthesis will be reported elsewhere. Lipids 1 and 2 differ by the length of the linker between the hydrophobic section and the NTA functional group. A longer linker was thought to give more freedom of movement and thus to be conducive to a better crystal packing of the protein Lipid 3 was used as a diluting lipid matrix. Lipid 4 is the regular non-¯uorinated lipid used for comparative studies. Monolayer properties of the lipids 1 and 2 were investigated by mechanical and optical techniques. Isothermal surface pressure versus area curves
Figure 2. Structure of the partially ¯uorinated Ni2chelating lipids used in this study. The lipids were designed to spread at the air-water interface into monolayers in the ¯uid LE (liquid expanded) phase. Lipids 1 and 2 differ by the length of the linker between the fatty chains and the NTA moiety. Lipid 3 was used as a diluting lipid in the crystallization experiments. Lipid 4 (Ni-NTA-DOGA) was used for comparative experiments.
641 were determined using a Langmuir trough equipped with an electrobalance. Pro®les indicate that the two ¯uorinated lipids form stable monolayers at the air-water interface (Figure 3). It is well known the lipid matrix must be spread in the ¯uid liquid-expanded (LE) phase for successful 2D crystallization of proteins on lipid monolayers (Lebeau et al., 1996). This is essential to allow in-plane motions of the lipid-protein complexes so they can diffuse laterally and possibly organize into 2D arrays. An adequate ¯uidity of the lipid layers is ensured by branched aliphatic chains, which prevent tight, liquid-crystalline packing (Menger et al., 1988). On the pressure versus area curves, a plateau is observed at 1-2 nm2/molecule that is interpreted as the result of a conformational change of the branched aliphatic chains during monolayer compression in the ¯uid phase. The stability of the ¯uorinated monolayers in the presence of detergents was monitored by ellipsometry, an optical technique which is especially suited to the study of liquid surfaces. It is a non-invasive method that is sensitive to the density and thickness of the interface layer (Berge & Renault, 1993). The technique has been used to investigate the adsorption of proteins onto lipid monolayers (Andree et al., 1990; Caide et al., 1996; VeÂnien-Bryan et al., 1998). The ellipsometric angle d correlates with the amount of lipid and detergent adsorbed at the interface and is taken to be zero for a pure air-water interface. Spreading the non-¯uorinated nickel-chelating lipid 4 (Ni-NTADOGA, Figure 2) at the interface to form a monolayer produces a shift in d to 21 (Figure 4(a), arrow 1). The monolayer proves to be perfectly stable. If detergent is added in the aqueous phase (Figure 4(a), arrow 2), the value for the ellipsometric angle decreases within seconds and the ellipsometric angle reaches d 2(0.5) . This ellipsometric angle represents the value obtained for a
Figure 3. Isothermal surface pressure vs. lipid molecular area curves obtained with lipids 1 (thin line) and 2 (bold line) spread at the air-water interface at 20 C. Experimental procedures are described elsewhere (Lebeau et al., 1988).
642
Figure 4. Effect of detergents on the lipid monolayers monitored by ellipsometry. (a) Monolayer made made of non-¯uorinated lipid 4 (Ni-NTA-DOGA): the lipid is spread into a monolayer at the interface (arrow 1), then detergent (Emulgen 911, 0.5 CMC) is added into the aqueous subphase (arrow 2). (b)-(g) Monolayers made of ¯uorinated lipid 2. The experiment in (b) is similar to that described in (a): the lipid is spread at the interface (arrow 1), then Emulgen 911 is sequentially added into the aqueous subphase (arrow 2: 0.5 CMC; arrow 3: 0.5 CMC; arrow 4: 0.5 CMC; arrow 5: 1.5 CMC; arrow 6: 2.0 CMC). Various detergents have been tested: (c) Emulgen 911; (d) dodecyl-b-D-maltoside; (e) CHAPS; (f) SDS; (g) octyl-b-D-glucopyranoside. Arrows indicate addition of detergent (0.5 CMC). The loose dashed line refers to the value of the ellipsometric angle d recorded after addition of the corresponding detergent with no ¯uorinated lipid spread at the interface. The close dashed line indicates the equilibrium value for d when no detergent is added.
Membrane Protein 2D Crystallization on Lipid Layers
turbation at the interface. After a few minutes however the system goes back to equilibrium and d stabilizes close to the initial value of 7 . Repeated additions of detergent (Figure 4(b): arrow 3: 0.5 CMC; arrow 4: 0.5 CMC; arrow 5: 1.5 CMC; arrow 6: 2.0 CMC) to a ®nal ®vefold CMC detergent concentration only transiently disturbs the monolayer. Therefore the detergent Emulgen 911 is unsuccessful at solubilizing the monolayer of lipid 2. The ellipsometric angle remains constant for detergent Emulgen 911 concentration up to 5 CMC over a period of days. The stability of the ¯uorinated monolayer was investigated with detergents that have an extended range of CMC values (Figure 4(c)-(g)). High CMC detergents (octyl-b-D-glucopyranoside, CMC 7.3 g/l; sodium dodecyl sulfate, CMC 2.3 g/l) do not destabilize the ¯uorinated matrix more than those with lower CMC values (Emulgen 911, CMC 0.2 g/l; dodecyl-b-Dmaltoside, CMC 0.08 g/l). Some minor deviations of the d value can be interpreted as partial accumulation of detergent molecules at the polar interface created by the head groups of the nickelchelating lipids. This effect is especially noticeable with the CHAPS detergent (Figure 4(e)). It is possible for the sulphonate moiety in this detergent molecule to displace one or both water molecules from the nickel head group of lipid 2. This would then allow one or two detergent molecules to be tethered at the interface by one nickel ion, modifying the ellipsometric signal. But in no case are the ¯uorinated lipids removed from the interface and solubilized by the detergents. Lipid 1 exhibits very similar properties (data not shown). In some cases we have noticed that the ellipsometric signal obtained with the ¯uorinated lipids is more noisy than the signal obtained with the non-¯uorinated lipids. This is particularly visible in Figure 4(b). These ¯uctuations are interpreted as the presence of some heterogeneous lipid domains moving at the air-water interface. It is interesting to note that these ¯uctuations disappear when detergent is added (Figure 4). We conclude from these experiments that the partially ¯uorinated lipids spread at the air-water interface are resistant to solubilization by a large variety of detergents. Formation of 2D crystals using the fluorinated lipid monolayers
water-detergent solution. This indicates that the addition of even a small quantity of detergent (0.5 CMC) to the subphase under the lipid monolayer results in the complete and rapid solubilization of the lipid at the interface. The same experiment with the ¯uorinated lipid 1 or 2 takes a totally different course. Deposition of the ¯uorinated lipid at the air-water interface increases the d value to 7(1) (Figure 4(b), arrow 1). The addition of Emulgen 911 (Figure 4(b) arrow 2: 0.5 CMC) in the aqueous subphase provokes a transient per-
In order to check that the newly designed lipids were good candidates for promoting the formation of protein 2D crystals, we ®rst tested the lipids with streptavidin. This soluble protein has been extensively studied and the crystallization conditions are well known (Blankenburg et al., 1989). In this experiment, the partially ¯uorinated lipid was functionalized with a biotinyl group which binds tightly to streptavidin, rather than with the Ni-NTA group. Crystals of streptavidin were obtained reproducibly (data not shown), providing evidence that these ¯uorinated lipids possess
Membrane Protein 2D Crystallization on Lipid Layers
the correct ¯uidity and mobility properties at the interface necessary for the formation of crystals. We then used the ¯uorinated lipids functionalized with Ni-NTA moiety in order to crystallize membrane proteins. The plasma membrane H-ATPase (AHA2) from Arabidopsis thaliana is a single-subunit integral membrane protein with a molecular mass of 104 kDa. It belongs to the family of P-type transport ATPases. These are widely distributed biological energy transducers that convert the free energy resulting from ATP hydrolysis into an electrochemical ion gradient across the membrane (KuÈhlbrandt et al., 1998). Two-dimensional crystals of the corresponding enzyme from the bread mold Neurospora crassa have been obtained recently by surface crystallization on a solid carbon ®lm support. Well-ordered 2D crystals of the dodecylb-D-maltoside protein complex were found to grow rapidly in the presence of polyethylene glycol by a procedure based on that developed for growing 3D crystals (Cyrklaff et al., 1995). Even though the protein was not embedded in a lipid bilayer, the 2D crystals proved useful for structure analysis to Ê resolution (Auer et al., 1998, 1999). Two-dimen8A sional of the Neurospora crassa H-ATPase by conventional reconstitution into lipid membranes has so far been unsuccessful. The plant plasma membrane ATPase from A. thaliana was expressed in yeast with a C-terminal His-tag and puri®ed by Ni2-chelating chromatography. With the ¯uorinated lipids spread at the air-water interface, large well-ordered 2D crystals of the H-ATPase formed reproducibly. Crystals were obtained by spreading a mixture of lipids 2 and 3 at a ratio of 1:6 (w/w) onto 50 ml of the crystallization solution containing the protein and membrane lipids, both solubilized in dodecylb-D-maltoside. The drop was left at room temperature for a period of 24 hours during which the protein adsorbed to the ¯uorinated lipid layer. After this time, detergent was removed by adding 8-10 Bio-Beads1, followed by incubation for another 24-hour period to enable the crystal lattice to form. The Ni-NTA head group caused the protein to concentrate and orient at the interface between ¯uorinated lipid and water even when using a dilute protein solution (0.15 mg/ml). When samples were transfered to electron microscopy grids before the detergent removal step, large noncrystalline sheets of protein were observed (data not shown). At this stage the protein has attached to the lipid layer, probably as tightly clustered single molecules or oligomers while still being surrounded by detergent. Thus the crystal lattice clearly does not form before removal of detergent and protein insertion into a lipid bilayer. Complementary control experiments conducted in the presence of 250 mM imidazole or 150 mM NiSO4 under otherwise identical conditions did not produce crystals or even sheets of disordered protein (data not shown). As expected, both
643 chemicals were effective inhibitors of protein binding. Imidazole is used routinely in Ni2-chelating chromatography release to bound protein from the columns. Low concentrations of NiSO4 were suf®cient to saturate the Ni2-chelating histidine tags on the protein so that it was no longer able to attach to the derivatized lipid. These observations provide conclusive evidences that attachment of the protein via its His-tag to the Ni-NTA-derivatized lipid was an essential prerequisite for the formation of large, single layer crystals. For electron microscopy the 2D crystals were picked up from the surface of the drop with a carbon coated copper grid. Crystals were either negatively stained or prepared for electron cryomicroscopy by blotting, plunging into liquid ethane and mounting in a cryo-transfer holder. Grids were examined in an electron microscope and micrographs were recorded for structure determination by image analysis. Large membrane sheets, typically measuring more than 10 mm across, were frequently found and consisted of either a single crystal lattice, or of a few lattices fused in plane, as revealed by electron microscopy of negatively stained crystals (Figure 5). Projection map and structure of the H-ATPase Electron micrographs of frozen-hydrated 2D crystals were selected for coherent, well-ordered
Figure 5. 2D crystals of the Arabidopsis thaliana plasma membrane H-ATPase grown on a layer of lipids 2 and 3 after removal of the detergent, dodecyl-b-D-maltoside, by adsorption to Bio-Beads1. Crystals were stained with 2 % (w/w) uranyl acetate as a contrasting agent. (a) Overview of a large crystalline area. (b) The rectangular crystal lattice is visible at higher magni®cation.
644
Membrane Protein 2D Crystallization on Lipid Layers
lattices by optical diffraction, digitized and further analyzed by image processing (Crowther et al., 1996; Henderson et al., 1986). The unit cell parÊ , b 139 A Ê ameters were calculated as: a 151 A and g 90 (Table 1). Phase comparisons indicated that the crystal symmetry was P22121. Upon correction of lattice distortions, amplitudes and phases Ê resolution (Figure 6(a)). The were obtained to 9 A projection map (Figure 6(b)) looks different from the corresponding map of the Neurospora enzyme Ê . While the latter at a comparable resolution of 8 A crystallizes as a hexamer, the plant ATPase apparently crystallizes as a dimer. Each monomer consists of three distinct areas of density of roughly equal size in projection whereas the Neurospora ATPase does not show such a clear division into three density regions. The two enzymes are closely related by homology of the polypeptide sequence except that the N and C termini have different lengths (Axelsen & Palmgren, 1998). The projection maps must re¯ect these structural differences, but in addition may represent different conformations of the enzyme. This would not be surprising as the comparison of the Neurospora H-ATPase with the related Ca2-ATPase from sarcoplasmic reticulum has shown a high degree of conformational ¯exibility in the large cytoplasmatic domains, apparently associated with the E1 to E2 transition in the ion pumping cycle (Stokes et al., 1999). Such conformational changes could easily account for the observed differences in the projection maps. The P22121 symmetry of the 2D crystals of the plant H-ATPase indicates that the up and down orientation of each dimer alternates with respect to the plane of the lipid monolayer. This suggests that only one half of the molecules make speci®c contacts through their His-tag with the Ni-NTA moiety of the lipid head groups. The other half packs into the crystal in the upside-down orientation, resulting in a head-to-tail arrangement of pairs of ATPase molecules related by the 2-fold screw axes in the plane of the membrane. Small vesicular 2D crystals of the plant proton ATPase expressed in Saccharomyces cerevisiae have
been obtained by conventional detergent dialysis in the presence of membrane lipids (T.J., M.G.P., J.D., W.K. et al., unpublished results). It is interesting that these crystals have the same unit cell dimensions and symmetry as the crystalline sheets forming on lipid monolayers described here. They remain however much smaller, with diameters rarely exceeding 1-2 mm. Electron micrographs of the ATPase vesicle crystals always show two superimposed crystal lattices, one from each layer, whereas the 2D crystals described herein are single sheets, with only one lattice visible in the electron microscope. The fact that both crystal types show the same packing implies that the ATPase is embedded in a lipid bilayer both in the vesicle crystals and in the large crystalline sheets reported here. We cannot rule out that the crystalline sheets are produced by fusion of pre-formed crystalline vesicles which attach via their His-tags to the monolayers of ¯uorinated lipids. We think this mechanism of crystal formation is unlikely however because the protein clearly attaches to the monolayer prior to lattice formation, resulting in large, unordered layers which are not observed under conditions which prevent protein attachment. General use of fluorinated lipids for the 2D crystallization of membrane proteins Although it may be possible to grow 2D crystals either by conventional lipid reconstitution or on monolayers of detergent-resistant lipids, our new approach clearly has a number of important advantages for structure determination of membrane proteins by electron crystallography. The technique is quick and reproducible, and requires only minimal amounts of protein and functionalized lipid (approximately 7.5 mg and 500 nmol respectively per trial). The 2D crystals we obtained are large, single layers, offering the potential of electron diffraction as a straightforward test of crystalline order and state of preservation. This will make it much easier to optimize crystallization
Table 1. Electron crystallographic data Plane group symmetry Unit cell dimensions Number of images and lattices Range of defocus Total number of observations Number of unique observations Ê Overall phase residual to 9 A Ê )a Resolution range (A >20.4 20.4-14.3 14.3-11.6 11.6-10.1 10.1-9.0
P22121 Ê ; b138.9(1.5) A Ê ; g90 a150.9(1) A 5 Ê 7400-9500 A 611 191 25.5 Number of unique reflections Phase residualb 39 13.2 40 23.4 38 29.8 36 30.3 38 31.4
a Phase residual of re¯ections in resolution ranges, showing the mean deviation of averaged phases from the theoretical value, 0 or 180 . b Random data would give a value of 45 .
645
Membrane Protein 2D Crystallization on Lipid Layers
conditions for obtaining large, highly ordered 2D crystals and to collect data by electron diffraction and image processing. Finally, the approach described here may be applied to any membrane protein that can be expressed with a His-tag, and thus offers the prospect of a much needed rational, conceptually simple and potentially general method for the 2D crystallization of membrane proteins.
Materials and Methods Lipids, detergents and protein Phospholipids of the highest purity available were purchased from Avanti Polar-Lipids. Dodecyl-b-D-maltoside was from Calbiochem and Bio-Beads1 SM2 (20-50 mesh) from Bio-Rad. Ni-NTA-DOGA was prepared as described by (VeÂnien-Bryan et al. (1997). The synthesis of lipids 1, 2, and 3 will be described in detail elsewhere. Lipid stock solutions were kept at 1 or 2 mg/ml in chloroform/methanol (1:1, v/v) at ÿ20 C under argon gas. His-tagged H-ATPase was over-expressed and puri®ed to homogeneity as described (Jahn et al., 2001). Ellipsometry
Figure 6. (a) Structure factors of a 2D crystal of the Arabidopsis plasma membrane H-ATPase. The image was obtained by electron cryo-microscopy and image processing of unstained 2D crystals. The size of the square and the number re¯ect the signal-to-noise ratio of the computed Fourier component (Henderson et al., 1986). The concentric rings indicate the zero positions of the contrast transfer function. The resolution at the edge Ê . (b) Projection map of the plasma of the plot is 9 A Ê resolution calculated from membrane H-ATPase at 9 A merged amplitudes and phases from ®ve independent crystals with P22121 symmetry. Solid lines indicate density above the mean. Negative contours are indicated by dotted lines. The unit cell contains four molecules and Ê , b 139 A Ê and g 90 . The parameters are a 151 A monomer boundary cannot be de®ned with certainty in the projection map but comparison with the projected structure of the Neurospora enzyme suggests the outlined shape of the monomer.
The ellipsometric measurements were carried out with a conventional ellipsometer using a He-Ne laser operating at 632.8 nm as described by Berge & Renault (1993) and VeÂnien-Bryan et al. (1998). After re¯ection at the surface of water, the beam is analysed. The phase difference between the vertical and horizontal beam polarizations and so the ellipsometric angle (d) are determined. The variation of the ellisometric angle is a relevant probe for changes occurring at the interface. It is proportional to the amount of lipid and detergent adsorbed at the interface. The sample cell is made of Te¯on1 and has a volume of 8 ml. Basically, a lipid monolayer was spread at the air-water interface. Once equilibrium was reached, detergent was injected into the subphase. Homogeneity of the solution was ensured using a peristaltic pump. When the surface tension was measured in the same trough, we used the Wilhelmy method using a 1-cmwide ®lter paper. For more precise measurements of the surface pressure versus lipid molecular area we used a Wilhelmy plate coupled to a linear transducer (KSV 9900 Surface Barostat, KSV, Finland) and a Langmuir Te¯on1 trough (165 mm 270 mm). Monolayers were spread from 0.5 mM lipid solutions. Volume ranging from 100 to 300 ml were delivered at several locations across the water surface with a Hamilton microsyringe. At least 15 minutes were allowed for the spreading solvent to evaporate before monolayer compression. The subphase consisted of puri®ed water (pH 5.35, Millipore1 ®ltration system, Barnstead, NANO pure 11) with a resistivity of 18.2 106 cm. Surface pressure versus molecular area curves were recorded at 22.0(0.2) C with a comÊ 2 per lipid molecule per minute. pression speed of 2-5 A All measurements were carried out in triplicate. Protein binding and crystallization The protein (150 mg/ml) and egg phosphatidylcholine/dodecyl-b-D-maltoside mixed micelles (lipid to protein ratio 1:3, w/w) were placed in a Te¯on1 well (6 mm diameter, 1 mm deep) in a buffer containing 20 % (w/w) glycerol, 20 mM Mes, 150 mM KCl and a ®nal
646 detergent concentration of about 0.1 % (w/w) dodecylb-D-maltoside for membrane reconstitution. The resulting drop (50 ml) was coated with a monolayer of ¯uorinated lipid by adding 1 ml of the working solution (lipids 2 and 3 at a ratio of 1:6, w/w, 500 mM in chloroform/hexane 1:1, v/v) to the surface with a Hamilton syringe. After a ®rst 24-hour incubation period at 20 C in a humid chamber, eight to ten Bio-Beads1 were added to the well through a submerged channel, to avoid any disturbance of the drop surface. Two-dimensional crystals were allowed to form for another 24 hours and were transferred onto electron microscopy grids for analysis. Addition of 1 mM EDTA and 0.25 mM DTT to the subphase to stabilize the protein did not interfere with the formation of crystalline monolayers. These conditions were similar to those used routinely for column chromatography of the His-tagged ATPase on Ni2-chelating columns (M.J., M.G.P., J.D., W.K. et al., unpublished results). Control experiments in the presence of 10 mM EDTA, 150 mM NiSO4 or 250 mM imidazole which would release the His-tagged protein from the NTA column did not yield any protein crystalline sheets. Electron microscopy and image analysis Carbon-coated electron microscope grids were baked at 150 C for one hour prior to use in order to enhance the adhesion of the crystals by making the grids more hydrophobic. The crystals were picked up by placing a carbon-coated grid on top of the crystallization well for several minutes. The grid was removed, blotted with ®lter paper and either negatively stained with 2 % (w/w) uranyl acetate for 30 seconds or, plunged into liquid ethane and transferred to a GATAN cryo-transfer stage. Images were recorded on a Philips CM 120 transmission electron microscope operating at 120 kV in the low dose mode at a magni®cation of 45,000 . Selected images were processed using the MRC program suite (Crowther et al., 1996; Henderson et al., 1986).
Acknowledgements We are grateful to Declan Doyle for his critical reading of the manuscript and constructive comments. We thank the AFM (France) for a grant to F.L. The EU Biotechnology Program is acknowledged for ®nancial support (M.G.P. and W.K.).
References Andree, H. A. M., Reutelingsperger, C. P. M., Hauptmann, R., Hemker, H. C., Hermens, W. T. & Willems, G. M. (1990). Binding of vascular anticoagulant a (VAC a) to planar phospholipid bilayers. J. Biol. Chem. 265, 4923-4928. Auer, M., Scarborough, G. A. & KuÈhlbrandt, W. (1998). Three-dimensional map of the plasma membrane H-ATPase in the open conformation. Nature, 392, 840-841. Auer, M., Scarborough, G. A. & KuÈhlbrandt, W. (1999). Surface crystallization of the plasma membrane H-ATPase on a carbon support ®lm for electron crystallography. J. Mol. Biol. 287, 961-968. Axelsen, K. B. & Palmgren, M. G. (1998). Evolution of substrate speci®cities in the P-type ATPase superfamily. J. Mol. Evol. 46, 84-101.
Membrane Protein 2D Crystallization on Lipid Layers Berge, B. & Renault, A. (1993). Ellipsometry study of 2D crystallization of 1-alcohol monolayers at the water interface. Europhys. Letters, 21, 773-777. Blankenburg, R., Meller, P., Ringsdorf, H. & Salesse, C. (1989). Interaction between biotin lipids and streptavidin in monolayers: formation of oriented twodimentionsal protein domains induced by surface recognition. Biochemistry, 28, 8214-8221. Caide, X., Liu, Z., Gao, Q. F., Zhou, Q. Z. & Sui, S. F. (1996). Speci®c interaction of rabbit C-reactive protein with phospholipid membranes. Thin Solid Films, 285, 793-796. Crowther, R. A., Henderson, R. & Smith, J. M. (1996). MRC Image processing programs. J. Struct. Biol. 116, 9-16. Cyrklaff, M., Auer, M., KuÈhlbrandt, W. & Scarborough, G. A. (1995). 2-D structure of Neurospora crassa plasma membrane ATPase as determined by electron cryomicroscopy. EMBO J. 14, 1854-1857. Dietrich, C., Schmitt, L. & TampeÂ, R. (1995). Molecular organization of histidine-tagged biomolecules at self-assembled lipid interfaces using a novel class of chelator lipids. Proc. Natl Acad. Sci. USA, 92, 90149018. Frishman, D. & Mewes, H. W. (1997). Protein structural classes in ®ve complete genomes. Nature Struct. Biol. 4, 626-628. Held, P., Lach, F., Lebeau, L. & Mioskowski, C. (1997). Synthesis and preliminary evaluation of a new class of ¯uorinated amphiphiles designed for in-plane immobilization of biological macromolecules. Tetrahedron Letters, 38, 1937-1940. Helenius, A. & Simons, K. (1975). Solubilization of membrane proteins by detergents. Biochim. Biophys. Acta, 415, 29-79. Henderson, R., Baldwin, J. M., Downing, K. H., Lepault, J. & Zemlin, F. (1986). Structure of purple membrane from Halobacterium halobium: recording, measurement and evaluation of electron microÊ resolution. Ultramicroscopy, 19, 147graphs at 3.5 A 178. Henderson, R., Baldwin, J. M., Ceska, T. A., Zemlin, F., Beckmann, E. & Downing, K. H. (1990). Model for the structure of bacteriorhodopsin based on highresolution electron cryomicroscopy. J. Mol. Biol. 213, 899-929. Hochuli, E., DoÈbeli, H. & Schacher, A. (1987). New metal chelate adsorbent selective for proteins and peptides containing neighbouring histidine residues. J. Chromatog. 411, 177-184. Jahn, T., Dietrich, J., Andersen, B., Leidvik, B., Otter, C., Briving, C., KuÈhlbrandt, W. & Palmgren, M. G. (2001). Large scale expression, puri®cation and 2D crystallization of recombinant plant plasma membrane HATPase. J. Mol. Biol. In the press. Jap, B. K., Zulauf, M., Scheybani, T., Hefti, A., Baumeister, W., Aebi, U. & Engel, A. (1992). 2D crystallization: from art to science. Ultramicroscopy, 46, 45-84. Jones, D. T. (1998). Do transmembrane protein superfolds exist? FEBS Letters, 423, 281-285. Kubalek, E. W., Le Grice, S. F. J. & Brown, P. O. (1994). Two-dimensional crystallization of histidine-tagged HIV-1 reverse transcriptase promoted by a novel nickel-chelating lipid. J. Struct. Biol. 113, 117-123. KuÈhlbrandt, W., Wang, D. N. & Fujiyoshi, Y. (1994). Atomic model of plant light-harvesting complex by electron crystallography. Nature, 367, 614-621.
Membrane Protein 2D Crystallization on Lipid Layers KuÈhlbrandt, W., Auer, M. & Scarborough, G. A. (1998). Structure of the P-type ATPases. Curr. Opin. Struct. Biol. 8, 510-516. Landau, E. M. & Rosenbusch, J. P. (1996). Lipidic cubic phases: a novel concept for the crystallization of membrane proteins. Proc. Natl Acad. Sci. USA, 93, 14532-14535. Lebeau, L., Mioskowski, C. & Oudet, P. (1988). Systematic study of phospholipids linked to a steroid derivative, spread into a monolayer at the air/ water interface. Biochim. Biophys. Acta, 939, 417-429. Lebeau, L., Schultz, P., CeÂlia, H., MeÂsini, P., Nuss, S. & Klinger, C. et al. (1996). Speci®cally designed lipid assemblies as tools for two-dimensional crystallization of soluble biological macromolecules. In Handbook of Non-medical Applications of Liposomes (Barenholz, Y. & Lasic, D. D., eds), vol. II pp. 153186, CRC Press, Boca Raton. LeÂvy, D., Mosser, G., Lambert, O., Moeck, G. S., Bald, D. & Rigaud, J. L. (1999). Two-dimensional crystallization on lipid layer: a successful approach for membrane proteins. J. Struct. Biol. 127, 44-52. Menger, F. M., Wood, M. G. J., Richardson, S., Zhou, Q., Elrington, A. R. & Sherrod, M. J. (1988). Chainsubstituted lipids in monolayer ®lms. A study of molecular packing. J. Am. Chem. Soc. 110, 6797-6803. Mukerjee, P. (1994). Fluorocarbon-hydrocarbon interactions in micelles and other lipid assemblies, at interfaces, and in solutions. Colloids Surf. A, 84, 1-10.
647 Mukerjee, P. & Yang, A. Y. S. (1976). Non-ideality of mixing of micelles of ¯uorocarbon and hydrocarbon surfactants and evidence of partial miscibility from differential conductance data. J. Phys. Chem. 80, 1388-1390. Stokes, D. L., Auer, M., Zhang, P. & KuÈhlbrandt, W. (1999). Comparison of H-ATPase and Ca2-ATPase suggests that a large conformational change initiates P-type ion pump reaction cycles. Curr. Biol. 9, 672679. Tanford, C. & Reynolds, J. A. (1976). Characterization of membrane proteins in detergent solutions. Biochim. Biophys. Acta, 457, 133-170. VeÂnien-Bryan, C., Balavoine, F., Toussaint, B., Mioskowski, C., Hewat, E. A., Helme, B. & Vignais, P. M. (1997). Structural study of the response regulator HupR from Rhodobacter capsulatus. Electron microscopy of two-dimensional crystals on a nickelchelating lipid. J. Mol. Biol. 274, 687-692. VeÂnien-Bryan, C., Lenne, P. F., Zakri, C., Renault, A., Brisson, A., Legrand, J. F. & Berge, B. (1998). Characterization of the growth of 2D protein crystals on a lipid monolayer by ellipsometry and rigidity measurements coupled to electron microscopy. Biophys. J. 74, 2649-2657. Walz, T. & Grigorieff, N. (1998). Electron crystallography of two-dimensional crystals of membrane proteins. J. Struct. Biol. 121, 142-161.
Edited by R. Huber (Received 8 November 2000; received in revised form 16 February 2001; accepted 12 March 2001)