Organization of the G Protein-coupled Receptors Rhodopsin and Opsin in Native Membranes
2003; Elsevier BV; Volume: 278; Issue: 24 Linguagem: Inglês
10.1074/jbc.m302536200
ISSN1083-351X
AutoresYan Liang, Dimitrios Fotiadis, Sławomir Filipek, David A. Saperstein, Krzysztof Palczewski, Andreas Engel,
Tópico(s)Retinal Development and Disorders
ResumoG protein-coupled receptors (GPCRs), which constitute the largest and structurally best conserved family of signaling molecules, are involved in virtually all physiological processes. Crystal structures are available only for the detergent-solubilized light receptor rhodopsin. In addition, this receptor is the only GPCR for which the presumed higher order oligomeric state in native membranes has been demonstrated (Fotiadis, D., Liang, Y., Filipek, S., Saperstein, D. A., Engel, A., and Palczewski, K. (2003) Nature 421, 127–128). Here, we have determined by atomic force microscopy the organization of rhodopsin in native membranes obtained from wild-type mouse photoreceptors and opsin isolated from photoreceptors of Rpe65–/– mutant mice, which do not produce the chromophore 11-cis-retinal. The higher order organization of rhodopsin was present irrespective of the support on which the membranes were adsorbed for imaging. Rhodopsin and opsin form structural dimers that are organized in paracrystalline arrays. The intradimeric contact is likely to involve helices IV and V, whereas contacts mainly between helices I and II and the cytoplasmic loop connecting helices V and VI facilitate the formation of rhodopsin dimer rows. Contacts between rows are on the extracellular side and involve helix I. This is the first semi-empirical model of a higher order structure of a GPCR in native membranes, and it has profound implications for the understanding of how this receptor interacts with partner proteins. G protein-coupled receptors (GPCRs), which constitute the largest and structurally best conserved family of signaling molecules, are involved in virtually all physiological processes. Crystal structures are available only for the detergent-solubilized light receptor rhodopsin. In addition, this receptor is the only GPCR for which the presumed higher order oligomeric state in native membranes has been demonstrated (Fotiadis, D., Liang, Y., Filipek, S., Saperstein, D. A., Engel, A., and Palczewski, K. (2003) Nature 421, 127–128). Here, we have determined by atomic force microscopy the organization of rhodopsin in native membranes obtained from wild-type mouse photoreceptors and opsin isolated from photoreceptors of Rpe65–/– mutant mice, which do not produce the chromophore 11-cis-retinal. The higher order organization of rhodopsin was present irrespective of the support on which the membranes were adsorbed for imaging. Rhodopsin and opsin form structural dimers that are organized in paracrystalline arrays. The intradimeric contact is likely to involve helices IV and V, whereas contacts mainly between helices I and II and the cytoplasmic loop connecting helices V and VI facilitate the formation of rhodopsin dimer rows. Contacts between rows are on the extracellular side and involve helix I. This is the first semi-empirical model of a higher order structure of a GPCR in native membranes, and it has profound implications for the understanding of how this receptor interacts with partner proteins. Vision is essential for the survival of many organisms ranging from unicellular dinoflagellates to man (1Gehring W.J. Int. J. Dev. Biol. 2002; 46: 65-73PubMed Google Scholar). Rhodopsin, the primary molecule in the visual signaling cascade, is activated by a single photon and induces subunit dissociation of transducin (Gt) 1The abbreviations used are: Gt, transducin (rod photoreceptor G-protein); AFM, atomic force microscope/microscopy; GPCR, G protein-coupled receptor; ROS, rod outer segment(s); EM, electron microscopy; RPE cells, retinal pigment epithelial cells. molecules, the cognate G proteins, amplifying the light signal (2Stryer L. Biopolymers. 1985; 24: 29-47Crossref PubMed Scopus (29) Google Scholar). Rhodopsin is also a prototypical G protein-coupled receptor (GPCR) and a member of subfamily A, which comprises ∼90% of all GPCRs (3Ballesteros J. Palczewski K. Curr. Opin. Drug Discov. Dev. 2001; 4: 561-574PubMed Google Scholar). GPCRs are essential proteins in signal transduction across cellular membranes (4Pierce K.L. Premont R.T. Lefkowitz R.J. Nat. Rev. Mol. Cell. Biol. 2002; 3: 639-650Crossref PubMed Scopus (2125) Google Scholar). The first crystal structure of a GPCR, rhodopsin, has been determined (5Palczewski K. Kumasaka T. Hori T. Behnke C.A. Motoshima H. Fox B.A. Le Trong I. Teller D.C. Okada T. Stenkamp R.E. Yamamoto M. Miyano M. Science. 2000; 289: 739-745Crossref PubMed Scopus (5056) Google Scholar), and two refined models have subsequently been reported (6Teller D.C. Okada T. Behnke C.A. Palczewski K. Stenkamp R.E. Biochemistry. 2001; 40: 7761-7772Crossref PubMed Scopus (629) Google Scholar, 7Okada T. Fujiyoshi Y. Silow M. Navarro J. Landau E.M. Shichida Y. Proc. Natl. Acad. Sci. U. S. A. 2002; 99: 5982-5987Crossref PubMed Scopus (660) Google Scholar). In vertebrate retinal photoreceptors, rod outer segment (ROS) disk membranes are tightly stacked (8Molday R.S. Invest. Ophthalmol. Vis. Sci. 1998; 39: 2491-2513PubMed Google Scholar). The stacking of these internal cellular membrane structures ensures a dense packing of light-absorbing rhodopsins, which constitute >90% of all disk membrane proteins, and in turn, a high probability of single photon absorption (9Baylor D.A. Lamb T.D. Yau K.W. J. Physiol. 1979; 288: 613-634PubMed Google Scholar). In ROS disk membranes, rhodopsin occupies ∼50% of the space within the disks (8Molday R.S. Invest. Ophthalmol. Vis. Sci. 1998; 39: 2491-2513PubMed Google Scholar). Knockout mice lacking rhodopsin do not develop ROS, which indicates a structural role for this protein (10Lem J. Krasnoperova N.V. Calvert P.D. Kosaras B. Cameron D.A. Nicolo M. Makino C.L. Sidman R.L. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 736-741Crossref PubMed Scopus (334) Google Scholar, 11Humphries M.M. Rancourt D. Farrar G.J. Kenna P. Hazel M. Bush R.A. Sieving P.A. Sheils D.M. McNally N. Creighton P. Erven A. Boros A. Gulya K. Capecchi M.R. Humphries P. Nat. Genet. 1997; 15: 216-219Crossref PubMed Scopus (459) Google Scholar). The organization of rhodopsin and other GPCRs in their native membranes is of paramount importance because the physiological properties of these receptors may depend on their oligomeric state (reviewed in Refs. 4Pierce K.L. Premont R.T. Lefkowitz R.J. Nat. Rev. Mol. Cell. Biol. 2002; 3: 639-650Crossref PubMed Scopus (2125) Google Scholar and 12George S.R. O'Dowd B.F. Lee S.P. Nat. Rev. Drug Discov. 2002; 1: 808-820Crossref PubMed Scopus (546) Google Scholar, 13Bouvier M. Nat. Rev. Neurosci. 2001; 2: 274-286Crossref PubMed Scopus (584) Google Scholar, 14Rios C.D. Jordan B.A. Gomes I. Devi L.A. Pharmacol. Ther. 2001; 92: 71-87Crossref PubMed Scopus (291) Google Scholar). In native disk membranes, the existence of distinct, densely packed rows of rhodopsin dimers has been demonstrated by AFM (15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar). To obtain further insight into the native molecular organization of GPCRs, we have used AFM to visualize the organization of rhodopsin and opsin in their native membranes. Finally, we produced a model of rhodopsin oligomers that accounts for all geometrical constraints imposed by AFM and crystallographic data. This model shows, for the first time, the higher order organization of a GPCR in its native environment. The contact sites identified in the model, which are responsible for the oligomerization of rhodopsin, are likely to be crucial for the self-assembly of other GPCRs (4Pierce K.L. Premont R.T. Lefkowitz R.J. Nat. Rev. Mol. Cell. Biol. 2002; 3: 639-650Crossref PubMed Scopus (2125) Google Scholar, 12George S.R. O'Dowd B.F. Lee S.P. Nat. Rev. Drug Discov. 2002; 1: 808-820Crossref PubMed Scopus (546) Google Scholar, 13Bouvier M. Nat. Rev. Neurosci. 2001; 2: 274-286Crossref PubMed Scopus (584) Google Scholar, 14Rios C.D. Jordan B.A. Gomes I. Devi L.A. Pharmacol. Ther. 2001; 92: 71-87Crossref PubMed Scopus (291) Google Scholar). Isolation of ROS and Disk Membranes—All animal experiments employed procedures approved by the University of Washington Animal Care Committee. Rpe65-deficient mice and wild-type C57BL/6 mice were obtained from M. Redmond (National Eye Institute) (16Redmond T.M. Yu S. Lee E. Bok D. Hamasaki D. Chen N. Goletz P. Ma J.X. Crouch R.K. Pfeifer K. Nat. Genet. 1998; 20: 344-351Crossref PubMed Scopus (797) Google Scholar) and The Jackson Laboratory, respectively. All animals (4–8 weeks old) were maintained in complete darkness for >120 min before they were sacrificed. The eyes were removed and the retinas isolated in complete darkness with the aid of night vision goggles (Lambda 9, ITT Industries). Twelve mouse retinas were placed in a tube with 120 μl of 8% OptiPrep (Nycomed, Oslo, Norway) in Ringer's buffer (130 mm NaCl, 3.6 mm KCl, 2.4 mm MgCl2, 1.2 mm CaCl2,10mm Hepes, pH 7.4, containing 0.02 mm EDTA) and vortexed for 1 min. The samples were centrifuged at 200 × g for 1 min, and the supernatant containing the ROS was removed gently. The pellet was dissolved in 120 μl of 8% OptiPrep, vortexed, and centrifuged again. The vortexing and sedimentation sequence was repeated six times. The collected ROS supernatants (∼1.5 ml) were combined, overlaid on a 10–30% continuous gradient of OptiPrep in Ringer's buffer, and centrifuged for 50 min at 26,500 × g. ROS were harvested as a second band (about two-thirds of the way from the top), diluted three times with Ringer's buffer, and centrifuged for 3 min at 500 × g to remove the cell nuclei. The supernatant containing ROS was transferred to a new tube and centrifuged for 30 min at 26,500 × g. The pelleted material contained pure, osmotically intact ROS. ROS were burst in 2 ml of 2 mm Tris-HCl, pH 7.4, at 0 °C for 15 h. Disks were overlaid on a 15–40% continuous gradient of OptiPrep in Ringer's buffer. The sample was centrifuged for 50 min at 26,500 × g, and the disks were collected from a faint band located about two-thirds of the way from the top of the gradient. The harvested intact disks were then diluted three times with Ringer's solution and pelleted for 30 min at 26,500 × g. SDS-PAGE and immunoblotting were performed as described previously (17Ohguro H. Chiba S. Igarashi Y. Matsumoto H. Akino T. Palczewski K. Proc. Natl. Acad. Sci. U. S. A. 1993; 90: 3241-3245Crossref PubMed Scopus (60) Google Scholar). Phosphorylation and Reduction Reactions—Phosphorylation of rhodopsin and reduction of all-trans-retinal were carried out as described previously (18Palczewski K. McDowell J.H. Hargrave P.A. J. Biol. Chem. 1988; 263: 14067-14073Abstract Full Text PDF PubMed Google Scholar). The indicated samples were sonicated for 30 s in a Bransonic 220 sonicator (Fisher). Atomic Force Microscopy—Washed disk membranes were adsorbed to mica in 2 mm Tris-HCl, pH 7.4, for 15–20 min and washed with 20 mm Tris-HCl, pH 7.8, 150 mm KCl, 25 mm MgCl2 (recording buffer). AFM experiments were performed using a Nanoscope Multimode microscope (Digital Instruments) equipped with an infrared laser head, a fluid cell, and oxide-sharpened silicon nitride cantilevers (OMCL-TR400PSA, Olympus), calibrated as described previously (19Müller D.J. Engel A. Biophys. J. 1997; 73: 1633-1644Abstract Full Text PDF PubMed Scopus (236) Google Scholar). Topographs were acquired in contact mode at minimal loading forces (≤100 piconewtons). Trace and retrace signals were recorded simultaneously at line frequencies ranging between 4.1 and 5.1 Hz. The power spectrum displayed in the inset in Fig. 4a was calculated with the SEMPER image processing system (20Saxton W.O. J. Struct. Biol. 1996; 116: 230-236Crossref PubMed Scopus (67) Google Scholar). Scanning EM—The retinas without retinal pigment epithelial (RPE) cells were fixed in 2.5% glutaraldehyde, 0.1 m cacodylate buffer, 2% sucrose, pH 7.4, for 6 h. Fixed disks were allowed to settle on the coated (1% poly-l-lysine-coated) coverslip and washed with water. All samples were washed in 0.1 m cacodylate buffer, 2% sucrose, fixed with 1% OsO4 in washing buffer, dehydrated with ethanol, dried using a critical point drying method, sputter-coated with a 5–10-nm thick gold layer, and analyzed employing a JSF-6300F or an XL SFEG scanning electron microscope (FEI Sirion, Philips). Light and Transmission EM—ROS and disks were fixed in 2.5% glutaraldehyde, 1% OsO4, 0.13 m sodium phosphate, pH 7.4, for 1 h, washed three times using EM rinsing buffer (0.13 m NaH2PO4, 0.05% MgCl2, pH 7.4) and collected by centrifugation at 16,000 × g for 3 min. ROS and disk pellets were suspended in molten 5% phosphate-buffered low-temperature gelling agarose solution, collected by centrifugation at 16,000 × g for 3 min, and cooled. The ROS and disk pellets were secondarily fixed with 1% OsO4 in 0.1 m phosphate buffer, pH 7.4, dehydrated with ethanol, and embedded in Eponate12 resin (Ted Pella, Inc., Redding, CA). Thin sections (1.0 μm) were cut, stained with 10% Richardson's blue solution, and subjected to light microscopy. Ultrathin sections (0.07 μm) were cut and stained with uranyl acetate and lead citrate solution. Samples were recorded with a Philips CM-10 EM. Electron Microscopy of Immunogold-labeled and Negatively Stained Disk Membranes—Isolated disks were adsorbed to carbon support films mounted on electron microscopy grids, blocked with 0.5% bovine serum albumin in 150 mm NaCl, 50 mm Tris, pH 7.4, and incubated for 1.5 h with 1D4 (C-terminal specificity, R. Molday) (21MacKenzie D. Arendt A. Hargrave P. McDowell J.H. Molday R.S. Biochemistry. 1984; 23: 6544-6549Crossref PubMed Scopus (181) Google Scholar), 4D2 (N-terminal specificity, R. Molday) (22Laird D.W. Molday R.S. Invest. Ophthalmol. Vis. Sci. 1988; 29: 419-428PubMed Google Scholar), C7 (C-terminal specificity, K. Palczewski), or B6-30N (N-terminal specificity, P. Hargrave) (23Adamus G. Zam Z.S. Arendt A. Palczewski K. McDowell J.H. Hargrave P.A. Vision Res. 1991; 31: 17-31Crossref PubMed Scopus (172) Google Scholar) anti-rhodopsin antibody at dilutions of 1:10, 1:10, 1:1000, and 1:10, respectively. A secondary antibody, goat anti-mouse IgG conjugated with 15 nm gold, was used at a dilution of 1:100. Antibody-labeled and unlabeled disk membranes were stained with Nano-W negative stain (Nanoprobes, Stony Brook, NY) or 0.75% uranyl acetate, respectively. Electron micrographs were recorded with a Philips CM-10 or a Hitachi H-7000 electron microscope. The power spectra displayed in Fig. 5 were calculated with the SEMPER image processing system (20Saxton W.O. J. Struct. Biol. 1996; 116: 230-236Crossref PubMed Scopus (67) Google Scholar). Modeling—A monomer of the rhodopsin crystal structure (Protein Data Bank code 1HZH) (6Teller D.C. Okada T. Behnke C.A. Palczewski K. Stenkamp R.E. Biochemistry. 2001; 40: 7761-7772Crossref PubMed Scopus (629) Google Scholar) was used to build an oligomeric model of rhodopsin in the lipid membrane. The loops not present in the rhodopsin crystal structure were created using the Modeler module (24Sali A. Potterton L. Yuan F. van Vlijmen H. Karplus M. Proteins. 1995; 23: 318-326Crossref PubMed Scopus (958) Google Scholar) of Insight II (Insight II, version 2000, Accelrys, San Diego). Verification of the created loops and of the whole structure was accomplished with the Profile-3D module (25Bowie J.U. Luthy R. Eisenberg D. Science. 1991; 253: 164-170Crossref PubMed Scopus (2443) Google Scholar) by evaluating the compatibility between sequence and structure. The MOLMOL program was used to analyze the modeled macromolecular structures (26Koradi R. Billeter M. Wuthrich K. J. Mol. Graph. 1996; 14 (:29–32): 51-55Crossref PubMed Scopus (6498) Google Scholar). This theoretical model of the native rhodopsin organization was deposited in the Protein Data Bank under the accession number 1N3M. Isolation and Characterization of ROS and Disk Membranes—ROS were isolated from mouse retinae. As demonstrated by transmission EM, scanning electron, and light microscopy, the protocol employed yielded highly enriched and structurally preserved ROS with a diameter of 0.85–1.4 μm and a length of 6–10 μm (Fig. 1). Thus, the diameters of ROS still attached to the retina (Fig. 1a) and isolated ROS (Fig. 1, b–d) were comparable, suggesting structural integrity. Moreover, to check the quality of our preparations, we employed UV-visible spectroscopy and enzymatic assays of rhodopsin phosphorylation using intracellular endogenous rhodopsin kinase (18Palczewski K. McDowell J.H. Hargrave P.A. J. Biol. Chem. 1988; 263: 14067-14073Abstract Full Text PDF PubMed Google Scholar) and reduction of the photoisomerized chromophore of rhodopsin (27Palczewski K. Jager S. Buczylko J. Crouch R.K. Bredberg D.L. Hofmann K.P. Asson-Batres M.A. Saari J.C. Biochemistry. 1994; 33: 13741-13750Crossref PubMed Scopus (133) Google Scholar), all-trans-retinal, using membrane impermeable [γ-32P]ATP and [C4-3H]NADPH, respectively (Fig. 1e). The phosphorylation level was low (8.40 pmol) in untreated samples. Samples that were irradiated by light and subsequently sonicated expressed low quantities of phosphorylated rhodopsin as well. In contrast, the phosphorylation was the highest (49.40 pmol) in samples that were sonicated during light irradiation. The amount of [3H]retinol was lowest in untreated samples and remained low in samples that were sonicated after light irradiation. In samples sonicated under light irradiation, the quantity of [3H]retinol was the highest (Fig. 1e). Thus, the results indicated that the isolated ROS were osmotically intact and that the rhodopsin molecules were fully active. Disks isolated after osmotic bursting of the ROS and prepared by thin sectioning appeared as vesicles in the EM, compatible with the high osmotic pressure expected to inflate the structurally preserved disks (Fig. 1f). Immunogold labeling of disks was performed using antibodies directed against the N-terminal (4D2 antibody) and C-terminal (1D4 antibody) ends of rhodopsin. More than 90% of the disks bound the C-terminal anti-rhodopsin antibody throughout the disk surface (Fig. 1g, arrows). Less than 10% were labeled around their rim (Fig. 1g, inset 1), suggesting that disrupted disks expose their extracellular surface. In agreement, about 10% of the disks were labeled when using the antibody directed against the rhodopsin N terminus (Fig. 1g, inset 2). Taken together, the antibody labeling experiments strongly support the structural preservation of the disk membranes during their isolation. SDS-PAGE (Fig. 1h) revealed that the disk preparation did not contain significant amounts of soluble proteins normally present in ROS and was enriched in rhodopsin (>95%). The latter finding was identified by immunoblotting using the 4D2 and C7 antibodies (data not shown). AFM Imaging of Rhodopsin in Native Disk Membranes—To unveil the native supramolecular arrangement of rhodopsin, isolated disk membranes were adsorbed to freshly cleaved mica and imaged by AFM in buffer solution. The AFM was equipped with an infrared laser to avoid the formation of opsin, the retinal-depleted form of rhodopsin (28McBee J.K. Palczewski K. Baehr W. Pepperberg D.R. Prog. Retin. Eye Res. 2001; 20: 469-529Crossref PubMed Scopus (317) Google Scholar). The morphology of an intact native disk adsorbed to mica is revealed in Fig. 2. Three different surface types are evident: the cytoplasmic side of the disk (type 1), co-isolated lipid (type 2), and mica (type 3). Bare lipid bilayers had a thickness of 3.7 ± 0.2 nm (n = 86) and an unstructured topography (Fig. 2, type 2). Compared with the topography of the lipid, the cytoplasmic surface (type 1) of the disk was highly corrugated, indicating the presence of densely packed proteins (see deflection image in Fig. 2b). Well adsorbed, single- and double-layered disk membranes had a thickness of 7–8 nm and 16–17 nm, respectively, a circular shape, and diameters between 0.9 and 1.5 μm. These disk diameters, determined by AFM, are in excellent agreement with those obtained from ROS by scanning electron microscopy (Fig. 1, a and c) and light and electron microscopy (Fig. 1, b–d). Open, spread-flattened disks adsorbed as round-shaped single-layered membranes to mica and exhibited four different surface types (Fig. 3). The first surface type (Fig. 3a, type 1) was characterized by a highly textured topography consisting of densely packed double rows of protrusions forming paracrystals (Fig. 3b). SDS-PAGE revealed that rhodopsin was present at a high concentration in such disk membrane preparations (Fig. 1h), suggesting that the visualized densely packed rows and paracrystals are related to this major protein. The second and third surface types were the same as in Fig. 2, i.e. lipid and mica. The fourth surface type (Fig. 3a, type 4, and Fig. 3c) had the same morphology as the first except that the paracrystals formed rafts of rhodopsin separated by lipid. At higher magnification, rhodopsin dimers from densely packed regions (Fig. 3b, broken ellipses) or raft-like cluster (Fig. 3c, broken ellipses) to break off the rows were seen, identifying them as the building blocks of the paracrystals. Occasionally, single rhodopsin monomers (Fig. 3b, arrowhead) were detected on such topographs. The packing density in surface type 1 areas ranged between 30,000 and 55,000 rhodopsin monomers/μm2 (15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar), similar to the packing density within the rhodopsin islands in surface type 4 areas (in Fig. 3c the packing density is about 34,000 rhodopsin monomers/μm2). Obviously, the overall packing density of rhodopsin measured by AFM on tightly packed regions (Fig. 3b) or within rhodopsin rafts (Fig. 3c) is higher than that measured by optical methods (29Liebman P.A. Entine G. Science. 1974; 185: 457-459Crossref PubMed Scopus (205) Google Scholar).Fig. 3Topography of an open, spread-flattened disk adsorbed to mica and imaged in buffer solution.a, height image of the open, spread-flattened disk. Four different surface types are evident: the cytoplasmic surface of the disk (types 1 and 4), lipid (type 2), and mica (type 3). The topographies of regions 1 (b) and 4 (c) at higher magnification reveal densely packed rows of rhodopsin dimers. Besides paracrystals, single rhodopsin dimers (broken ellipses) and occasional rhodopsin monomers (arrowhead) are discerned floating in the lipid bilayer. Scale bars: 250 nm (a) and 15 nm (b and c). Vertical brightness ranges: 22 nm (a) and 2.0 nm (b and c).View Large Image Figure ViewerDownload Hi-res image Download (PPT) AFM Imaging of Opsin in Native Disk Membranes—The 65-kDa protein RPE65 is highly expressed in RPE cells and is one of the proteins involved in retinoid processing (reviewed in Ref. 28McBee J.K. Palczewski K. Baehr W. Pepperberg D.R. Prog. Retin. Eye Res. 2001; 20: 469-529Crossref PubMed Scopus (317) Google Scholar). In Rpe65–/– mice, retinoid analyses revealed no detectable 11-cis-products in any of the ester, aldehyde, or alcohol forms (16Redmond T.M. Yu S. Lee E. Bok D. Hamasaki D. Chen N. Goletz P. Ma J.X. Crouch R.K. Pfeifer K. Nat. Genet. 1998; 20: 344-351Crossref PubMed Scopus (797) Google Scholar). Although these mice are able to develop ROS, the ROS contain opsin instead of rhodopsin. We used preparations of disks from Rpe65–/– mice to compare the structure and the native supramolecular arrangement of opsin with that of rhodopsin. In general, the morphology of the Rpe65–/– disk membranes was similar to that of the wild-type membranes (see Fig. 3), but occasionally, even better ordered paracrystals could be found in Rpe65–/– preparations (Fig. 4a). From such areas, power spectra (Fig. 4a, inset) were calculated and the unit cell parameters determined (a = 8.4 ± 0.3 nm, b = 3.8 ± 0.2 nm, γ = 85 ± 2° (n = 9)), these values being the same as those found for wild-type paracrystals (15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar). At higher magnification, rows of opsin dimers forming the paracrystal (Fig. 4b, broken ellipse) were visualized, indicating the same oligomeric state as rhodopsin in its native environment (compare Fig. 4b with Fig. 3, b and c, and with Ref. 15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar). Occasionally, single-opsin monomers (Fig. 4b, arrowheads) were seen in such topographs. As with rhodopsin (15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar), opsin protruded by 1.4 ± 0.2 nm (n = 32) out of the lipid moiety on the cytoplasmic surface. On the extracellular surface, no opsin paracrystals were evident (Fig. 4c). The surface was corrugated, irregular, and flexible, preventing the acquisition of highly resolved AFM topographs such as required to reveal the paracrystalline packing. Opsin clusters (Fig. 4c, triangle) protruded 2.8 ± 0.2 nm (n = 60) out of the lipid bilayer (Fig. 4c, asterisk) on the extracellular side, which is twice the height of the cytoplasmic protrusions. The latter finding is also in line with the atomic structure of rhodopsin determined by x-ray crystallography (6Teller D.C. Okada T. Behnke C.A. Palczewski K. Stenkamp R.E. Biochemistry. 2001; 40: 7761-7772Crossref PubMed Scopus (629) Google Scholar). Irregular and flexible surfaces are typical for glycosylated proteins and proteins with long, flexible termini or loops. This observation, along with the fact that opsin has a long N terminus and is glycosylated on the extracellular surface, strengthens the assignment of this surface as the extracellular side of disk membranes. Similar difficulties were encountered with the glycosylated aquaporin-1 and the His-tagged AqpZ proteins where oligosaccharides or long termini impeded the acquisition of highly resolved surface topographs by AFM (30Scheuring S. Ringler P. Borgnia M. Stahlberg H. Muller D.J. Agre P. Engel A. EMBO J. 1999; 18: 4981-4987Crossref PubMed Scopus (165) Google Scholar, 31Walz T. Tittmann P. Fuchs K.H. Muller D.J. Smith B.L. Agre P. Gross H. Engel A. J. Mol. Biol. 1996; 264: 907-918Crossref PubMed Scopus (91) Google Scholar). Similar observations were also made for the membranes containing rhodopsin instead of opsin (data not shown). EM of Native Disk Membranes—To exclude the possibility of rhodopsin paracrystal formation upon adsorption on mica, native disk membranes were adsorbed on carbon-coated electron microscopy grids, negatively stained, and investigated by EM (Fig. 5). Power spectra were calculated from different regions of the adsorbed disks. Both the power spectra from a circular region adsorbed directly to the carbon film (Fig. 5, left PS) and from another region lying on disk membranes (Fig. 5, right PS) indicated diffraction patterns documenting the crystallinity of the disks irrespective of the support. Higher Order Organization of Rhodopsin in Native Membranes—The topographic information from AFM suggests a different packing arrangement of native rhodopsin dimers than of dimers observed in the three-dimensional crystal (5Palczewski K. Kumasaka T. Hori T. Behnke C.A. Motoshima H. Fox B.A. Le Trong I. Teller D.C. Okada T. Stenkamp R.E. Yamamoto M. Miyano M. Science. 2000; 289: 739-745Crossref PubMed Scopus (5056) Google Scholar). The thickness of single-layered disk membranes, 7.8 nm (15Fotiadis D. Liang Y. Filipek S. Saperstein D.A. Engel A. Palczewski K. Nature. 2003; 421: 127-128Crossref PubMed Scopus (661) Google Scholar), is compatible with the long axis, 7.5 nm, of the rhodopsin envelope derived from the 2.8-Å x-ray structure (6Teller D.C. Okada T. Behnke C.A. Palczewski K. Stenkamp R.E. Biochemistry. 2001; 40: 7761-7772Crossref PubMed Scopus (629) Google Scholar). This indicates that all rhodopsin molecules are integrated with their long axes perpendicular to the bilayer. The extracellular protrusion measured by AFM, 2.8 nm, is compatible with that estimated from the x-ray structure, 2.7 nm. However, the measured cytoplasmic protrusions of rhodopsin and of opsin, 1.4 nm, are significantly smaller than the 1.8 nm estimated from the atomic model. Unit cell dimensions (a = 8.4 nm, b = 3.8 nm, γ = 85°) of native rhodopsin and opsin paracrystals impose stringent boundary conditions for the packing arrangement of the rhodopsin/opsin dimers. The corresponding surface area (31.8 nm2) barely suffices to house two rhodopsin molecules whose cross-section fits in a rectangle of 4.8 × 3.7 nm2 (6Teller D.C. Okada T. Behnke C.A. Palczewski K. Stenkamp R.E. Biochemistry. 2001; 40: 7761-7772Crossref PubMed Scopus (629) Google Scholar). Thus, a small number of packing models emerged that were thoroughly tested for steric clashes and natures of contacts. The best model revealing the different intra- and interdimeric contacts is shown in Fig. 6a. The largest area of contact is 578 Å2 and intuitively represents the strongest interaction between rhodopsin molecules. It is found between helices IV and V, indicating this as the intradimeric contact (Fig. 6a, contact 1). Contacts involving helices I and II and the cytoplasmic loop between helices V and VI exhibit an area of 333 Å2 (Fig. 6a, contact 2) and represent the intra-row contacts. Finally, rows are weakly held together by interactions between regions of helix I close to the extracellular surface (Fig. 6a, contact 3) with a contact area of 146 Å2. Interactions within the Rho1–Rho2 dimer structure are located on both the cytoplasmic and the extracellular side. In the cytoplasmic part, hydrogen
Referência(s)