Detergent-Free Incorporation of a Seven-Transmembrane Receptor

Jul 24, 2012 - EaStCHEM School of Chemistry, University of Edinburgh, EH9 3JJ, United Kingdom. § Department of Chemistry, University of ..... Nano-si...
0 downloads 7 Views 292KB Size
Download PDF
Letter pubs.acs.org/NanoLett

Detergent-Free Incorporation of a Seven-Transmembrane Receptor Protein into Nanosized Bilayer Lipodisq Particles for Functional and Biophysical Studies Marcella Orwick-Rydmark,† Janet E. Lovett,‡ Andrea Graziadei,† Ljubica Lindholm,†,∥ Matthew R. Hicks,§,⊥ and Anthony Watts*,† †

Department of Biochemistry, Biomembrane Structure Unit, University of Oxford, Oxford, OX1 3QU, United Kingdom EaStCHEM School of Chemistry, University of Edinburgh, EH9 3JJ, United Kingdom § Department of Chemistry, University of Warwick, Library Road, Coventry, CV4 7AL, United Kingdom ‡

S Supporting Information *

ABSTRACT: SMA-Lipodisq nanoparticles, with one bacteriorhodopsin (bR) per 12 nm particle on average (protein/lipid molar ratio, 1:172), were prepared without the use of detergents. Using pulsed and continuous wave nitroxide spin label electron paramagnetic resonance, the structural and dynamic integrity of bR was retained when compared with data for bR obtained in the native membrane and in detergents and then with crystal data. This indicates the potential of Lipodisq nanoparticles as a useful membrane mimetic.

KEYWORDS: Lipodisq, bilayer, electron paramagnetic resonance, double electron electron resonance, bacteriorhodopsin, photoreceptor

M

(MSP), have the advantage that they can be made from a variety of lipid mixtures and have been shown to be relatively monodisperse in size upon protein incorporation.7,12 A drawback of nanodiscs is that they require detergents for protein incorporation, which must subsequently be completely removed for the assembly of a protein−nanodisc complex. Further, the absorbance properties of the membrane scaffold protein itself may interfere with the incorporated protein of interest or show specific lipid interactions with the rim protein. Lipodisq nanoparticles are lipid−polymer complexes that can be formed by detergent-free methods from a range of lipid compositions, and the polymer does not have the same interfering absorbance properties that nanodisc rim proteins possess.13−16 The application of the Lipodisq system composed of the lipid 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) and a polymer formed from a 3:1 mol:mol ratio of styrene to maleic acid, for the incorporation of the prototypical 7TM membrane protein, bacteriorhodopsin (bR), and subsequent biophysical applications are investigated here. Two distinguishable populations are present in a representative size exclusion chromatography (SEC) purification profile of bR-Lipodisq nanoparticles (Figure 1A). Peak 1 has a maximum at 565 nm, suggesting that this population is mainly composed of the purple membrane (PM) (Figure 1B), while peak 2

embrane proteins, upon removal from their native cell bilayer, should ideally be retained in a suitable hydrophobic environment that can maintain both protein functionality and structure. Detergent micelles are the most common medium for the initial removal of membrane proteins from the cell membrane and are often used for biophysical studies.1,2 However, detergent micelles are not necessarily a good mimetic for the membrane bilayer and may lead to protein aggregation, increased dynamics or (partial) unfolding, and the loss of structural integrity and hence activity.1,3 Membrane proteins have therefore been incorporated into nonmicellar systems such as liposomes,4 bicelles5,6 and more recently nanodiscs.7 While membrane proteins are often incorporated in liposomes, the resulting proteoliposomes are typically heterogeneously sized, with an inaccessible vesicle interior, resulting in difficulties in the analysis of functional studies requiring the addition of ligands or other cofactors to both sides of the membrane protein.8 Further, to maintain functionality, high protein−lipid ratios for biophysical studies are commonly required.9,10 Bicelles and nanodiscs are two alternative systems that provide accessibility to both faces of an incorporated protein and have proven useful for biophysical and structural studies on membrane proteins.8 However, bicelles can only be formed from very specific lipid compositions and can be heterogeneous in both size and composition.8,11 Apo-lipoprotein nanodiscs, which are composed of a lipid core surrounded by an amphipathic membrane scaffold protein © 2012 American Chemical Society

Received: May 30, 2012 Revised: July 23, 2012 Published: July 24, 2012 4687

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692

Nano Letters

Letter

Figure 1. (A) Size exclusion chromatography was used for the purification of bR-Lipodisq nanoparticles, with the absorbance is measured at 280 nm. (B) UV−visible spectra for Peak 1 in A, with Amax at 565 nm (red) and for Peak 2 with Amax at 555 nm (black). (C) DLS data for Peak 2 in a with a measured diameter of 12 ± 2 nm.

Figure 2 presents the CD spectra recorded for bR in the PM, reconstituted into DMPC vesicles, and incorporated into

possesses a maximal absorbance at 555 nm, indicating that bR is monomeric and incorporated into Lipodisq nanoparticles.17 Fractions from Peak 2 of the SEC purification protocol (Figure 1A) were analyzed by dynamic light scattering (DLS) to give a calculated diameter of 12 nm, 3 nm larger than the reported size of empty Lipodisq nanoparticles (Figure 1C) measured by us and others,13,14 although the extra membrane mass of bR may affect the diffusion coefficient of bR-Lipodisq nanoparticle, resulting in a slightly larger size.18 Assuming that an empty Lipodisq() nanoparticle contains between 180 and 200 lipids total,16 a bR monomer is likely to displace approximately 40 of the lipid molecules. Previously, a more homogeneous purification of bR-Lipodisq nanoparticles has been shown compared to that presented here.13 However, the previous SEC purification used a column one-third of the height used here (30 cm compared to 100 cm), such that sample inhomogeneities in that earlier study may have not been resolved. The molecule of retinal in bR is optically active in the presence of circularly polarized light, so that the visible circular dichroism (CD) spectrum of the bR retinal serves as an indicator for the oligomeric state of bR.19 The visible CD spectrum of bR in the PM is characterized by a biphasic band centered near the chromophore absorbance maximum which is attributed to the π−π* transition of the retinylidene group in the chromophore, due to the well-ordered crystalline nature of the PM and trimeric state of bR. Upon monomerization of bR, the biphasic band observed in the PM is reduced to a monophasic CD band.20 The reconstitution of bR into DMPC vesicles has been well-characterized by both electron microscopy and circular dichroism, and conditions such as the temperature and the ratio of bR:DMPC can be varied to modulate whether the trimeric or monomeric species of bR is formed.21 Previous studies confirm that below the DMPC main transition temperature at 23 °C22,23 and at high PM:DMPC lipid ratios (greater than ∼1:4 w/w) bR is able to form trimers in bilayers, while decreasing the PM/DMPC lipid ratio results in monomeric bR.21 Later studies revealed that the presence of 4 M NaCl and PM lipids could also induce trimeric bR independently of temperature.24,25 Therefore, the protocol used for the incorporation of bR into Lipodisq nanoparticles was carefully controlled so that the protein/total lipid molar ratio was 1:4 w/w, and the NaCl concentration was kept below 4 M (300 mM).

Figure 2. Visible CD spectra collected for bR in the PM (red line), in Lipodisq nanoparticles (black line), and reconstituted into DMPC vesicles (green line). The spectra represent the average of 60 scans and were recorded at 18 °C. The wavelength at maximum absorbance for each spectrum is noted.

Lipodisq nanoparticles at 18 °C, which is below the main phase liquid−gel DMPC transition temperature for protein-free bilayers (∼23 °C).22,23 These observations are consistent with bR monomerization upon incorporation into DMPC, which is then maintained upon addition of polymer to form bR-Lipodisq nanoparticles. The Lipodisq CD spectrum could not be simulated from any sum of the PM and DMPC spectra and is, therefore, not a mixture of the two states present in PM and DMPC (data not shown). Previously reported CD data for bR that is suggested to be incorporated into Lipodisq nanoparticles13 are more in agreement with that reported for bR reconstituted into DMPC at a similar protein/lipid molar ratio at 18 °C26 and is also consistent with CD data presented here for bR-DMPC proteoliposomes (Figure 2). The addition of the polymer results in an absorbance shift of 26 nm upfield, from 526 to 552 nm. bR D38C/F156C labeled with the commonly used nitroxide spin label, S-(1-oxy-2,2,5,5,-tetramethylpyrroline-3-methyl)methanethiosulfonate (MTSL) to give bR D38R1/F156R1,27 was used for all continuous wave (cw) and pulsed EPR measurements. An important question when working with spinlabeled oligomeric membrane proteins is whether the MTSL group is interacting with neighboring lipids. Indeed, previous 4688

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692

Nano Letters

Letter

provides only an approximate correlation time for the labeled system, it is useful to determine relative mobilities and changes in the spin label motion. For further quantification, a simple relationship between the reciprocal of the line-width of the central line of a nitroxide spectrum, ΔH0−1, which increases with increasing mobility of a spin label, can be correlated to the protein secondary structure36,37 and can be correlated to buried residues, tertiary contacts with other biomolecules, the helix/ surface interface, and to the loop region. The calculated τc for bR labeled at positions D38R1 and F156R1, located on the intracellular face of helices 2 and 5, respectively, in the PM and incorporated into Lipodisq nanoparticles are 10 and 8 ns, respectively, at RT, while solubilized bR undergoes considerably faster motional dynamics (5 ns; Table 1). These values indicate that the local

EPR studies with rhodopsin in both liposomes and detergents suggested that, depending on the position of the spin-label, interactions with the lipid headgroups could significantly influence the EPR spectra.28 Therefore, we used spin-labeling sites at the end of transmembrane (TM) helices 2 and 5, so that previous structural studies indicate do not interact with the lipid headgroups in the PM.29,30 Thus, we can infer that the continuous wave (cw) EPR spectra reflect bR backbone dynamics. In addition, particle tumbling is too slow (τc ∼100 ns) to average the nitroxide anisotropy.16 The cw EPR data for bR in the PM, in Lipodisq nanoparticles, and solubilized into octylglucoside (OG) detergent micelles are shown in Figure 3. The data in the

Table 1. τc and ΔH0−1 for bR D38R1/F156R1 in the PM, Lipodisq Nanoparticles, and OG Micelles. See Text for Estimation of τc and ΔH0−1 τc (ns)

ΔH0−1 (gauss−1)

PM Lipodisq nanoparticle OG micelles

10 ± 2 8±2 5±1

0.12 ± 0.01 0.15 ± 0.02 0.22 ± 0.04

dynamics at these labeled positions of the bR backbone are well-conserved within Lipodisq nanoparticles compared to the same position in the PM. The similarity in motional dynamics is confirmed by the values for (ΔH0)−1, which are again close for bR in the PM and in Lipodisq nanoparticles, at 0.12 and 0.15 G−1 (Table 1), respectively. In contrast, solubilization into OG increases the spin-label local motional freedom by approximately 50% (Table 1). Therefore, while previous structural studies indicate that OG is able to preserve the structural integrity of bR for crystallization,38 OG micelles do not faithfully maintain the backbone dynamics of bR at the monitored positions, in contrast to Lipodisq nanoparticles. Double-electron−electron resonance (DEER) spectroscopy is a pulsed EPR technique that allows for interspin distance measurements between 1.5 and 8 nm.39 The DEER signal is composed of both the intramolecular dipolar interaction between the two labels of interest, as well as the intermolecular dipolar interactions that result in an exponential decay referred to as the background. The background signal can be separated from the intramolecular dipolar interaction and the distance distribution between the labels of interest calculated.40 This technique has great potential for providing distance constraints within membrane proteins, as there are no size limitations on the biomolecule of interest and no need for long-range order.41 The phase memory, Tm, of a biomolecule is an important consideration for pulsed EPR measurements, as rapid signal decay can limit the interspin distance that can be determined and also leads to low signal-to-noise. For reconstituted membrane proteins, this is a particularly significant factor as reconstitution into a membrane environment results in a twodimensional background when compared to the three-dimensional background that is present in the bulk solvent, resulting in a high degree of spin diffusion between spin-labeled biomolecules and a short Tm.41 This effect can be mitigated by dilution with nonlabeled protein, though this requires considerably more protein, and decreases the total concentration of spin density such that measurements at standard Xband frequencies (∼ 9.4 GHz) become more difficult.42

Figure 3. bR D38R1/F156R1 cw EPR spectra in the native membrane (top), in Lipodisq nanoparticles (middle), and solubilized into OG micelles (bottom). The relatively more rigid (RR′) and less rigid (MM′) spectral components are expanded in the high field region. Dashed lines are at the motionally sensitive spectral extrema, 2A′zz, for the two component spectra, RR′ and MM′, and the solid lines mark the line width of the central peak, ΔH0.

PM display the composite spectra reported in previously published data for the single F156CR1 and D38R1 spectra in the PM31,32 and show that both the relative populations and positions of the mobile and less mobile components of the double mutant are remarkably well-preserved here in the Lipodisq nanoparticles, with subtle dynamic differences present compared to when bR is in the PM. The mobile component appears to be slightly more pronounced in the Lipodisq sample, which may be due to the shorter chain length of DMPC compared to native PM lipids.33 The detergent-solubilized sample is considerably more mobile than either the samples in the PM or in Lipodisq nanoparticles, indicating greater motional movement of the protein backbone when in detergent. This is likely because the detergent micelles are not able to stabilize bR dynamics as well as the Lipodisq nanoparticles can or because of partial solubilization and hence some unfolding of at least helices 2 and 5. An estimation of the correlation time for slow motional EPR spectra (τc > 10−9 s) can be achieved by: R b τc = a(A′zz − Azz )

sample preparation

(1)

2ARzz

where 2A′zz and are determined from the slow motion and rigid limit spectra, respectively.34,35 Although this approach 4689

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692

Nano Letters

Letter

Tm measurements were recorded for the bR double mutant D38R1/F156R1 in the PM, solubilized in OG, and incorporated into Lipodisq nanoparticles. A Tm of 400 ns was recorded in the PM, similar to that previously reported for reconstituted membrane proteins,42 suggesting spin diffusion that is likely due to both intramonomer and intertrimer spinlabel interactions. However, incorporation into Lipodisq nanoparticles or solubilizing the protein into detergent increased Tm 4-fold to 1400 and 1600 ns, respectively, similar to values reported for reconstituted membrane proteins that were spin-diluted.42 We can therefore infer that Lipodisq nanoparticles successfully isolate spin pairs from neighboring spin-labeled molecules, consistent with the CD data showing monomeric bR in Lipodisq nanoparticles (Figure 2). DEER distance measurements for bR D38R1/F156R1 in the PM, OG micelles, and Lipodisq nanoparticles show different dipolar evolution and distance distribution profiles (Figure 4).

samples of doubly labeled and spin-diluted for bR D38R1/ F156R1 in Lipodisq nanoparticles was also carried out.44,45 The results confirm the presence of weak dipolar coupling (Supporting Information, Figure 1), with a calculated interspin distance of 1.53 nm, in excellent agreement with the DEER data. Based upon the DEER and cw dipolar distance data, spinlabel distances were modeled into the bR crystal structure using the program Multiscale Modeling of Macromolecular systems (MMM2010; Figure 5).46

Figure 5. Nitroxide spin-labels attached to bR (PDB 1C3W), shown as balls and sticks at positions D38R1 (blue) and F156R1 (purple), with an interspin distance of 1.56 nm based upon DEER measurements.

Here we evaluate the ability of Lipodisq nanoparticles to maintain the dynamic and structural integrity of bR in a lipid environment. DLS measurements indicate an average bRLipodisq diameter of 12 nm (Figure 1C), and CD spectroscopy (Figure 2) confirms the monomeric form of the protein. cw EPR measurements (Figure 3) reveal that Lipodisq nanoparticles maintain the native bR dynamic profile observed in the PM, despite the increased lipid ordering observed in previous studies of Lipodisq nanoparticles16 which may have indicated that proteins incorporated into Lipodisq nanoparticles would have a motionally restricted profile compared to proteins in liposomes and other bilayer systems. Pulsed EPR results with bR incorporated into Lipodisq nanoparticles are consistent with bR monomerization and show that they are a suitable medium for DEER at X-band frequencies. Previously, where nanodiscs were used for DEER measurements on membrane proteins, the authors indicate difficulties in concentrating their samples such that Q-Band (34 GHz) was needed to measure the samples.47 This problem was not encountered when working with Lipodisq nanoparticles. Some 18 structures of bR have been reported in the ground state, largely by X-ray crystallography using detergentsolubilized bR, leading to inconsistencies at the loop-helical interface between the different structures.48 Lipodisq particles are one medium by which membrane proteins may be studied

Figure 4. Background-subtracted DEER data (fit shown in red) and the distance distribution obtained by Tikhonov regularization for bR D38R1/F156R1 in the PM (top panel), incorporated into Lipodisq nanoparticles (middle panel), and in OG micelles (bottom panel). Previous work suggests that, for short distances, a shallow modulation depth due to under-represented spin species can be observed, similar to that observed here.43

While data collected in the PM result in a time trace with no clear oscillation and several potential distances upon analysis (Figure 4, top panel), incorporation into Lipodisq nanoparticles results in a well-resolved oscillation and a distance distribution centered around 1.56 nm (Figure 4, middle panel), in good agreement with the Cα−Cα distance predicted from the crystal structure (Protein Data Bank ID 1C3W). This distance was also compared with the OG-solubilized sample, with a distance distribution centered at 1.58 nm (Figure 4, bottom panel). Compared to the Lipodisq sample, however, the oscillation is less pronounced in data for OG-solubilized bR, with a slightly broader distance distribution. As the DEER measurements here are on the borderline of applicability between cw and pulsed EPR distance measurements,43 a second moment analysis of the spectra for frozen 4690

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692

Nano Letters

Letter

integration time, scanning speed, and bandwidth were 1 s, 100 nm/min, and 2 nm, respectively. cw EPR measurements were measured on a cw Bruker EMX EPR spectrometer at RT. Samples were concentrated as above to approximately 2 mg/mL protein (76 μM) and loaded into 100 μL capillaries. Pulsed EPR measurements were carried out on a Bruker Elexsys 680 at X-Band using a 3 mm split ring resonator. Samples were concentrated to 200−300 μM. Phase memory measurements were recorded at 80 K. The four-pulse DEER experiment52 was employed ((π/2)v1 − τ1 − (π)v1 − t′(π)v2 − τ1 + τ2 − t′ − (π)v1 − τ2 − echo) with π/2 and π set to 32 ns, and the pump pulse equal to 12 ns. t′ was set to 80 ns and incremented by 8 ns. The first pulse was phase-cycled in the / channels, and the two signals were subtracted. The pump frequency was set to the central resonance, and the observer frequency was set 65 MHz upfield. Accumulation times were 16 h and measured at 80 K to avoid saturation of the signal. The data were processed with DeerAnalysis201153 using an experimental background dimensionality determined from a sparsely labeled sample prepared from a 3:1 molar ratio of the MTSL diagmagnetic analogue and MTSL. Rotameric libraries of R1 were generated and analyzed using MMM 2010.46

in the future without the need for detstabilizing detergents or perturbations induced by crystal contacts. Experimental Section. MTSL and its diamagnetic analogue (1-acetoxy-2,2,5,5-tetramethyl-δ-3-pyrroline-3methyl)methanethiosulfonate) were purchased from Toronto Research Chemicals (Ontario, Canada), and DMPC was purchased from Avanti Polar Lipids. The 3:1 SMA polymer (approximately MW = 9500 kDa) was supplied by Malvern Cosmeceutics Ltd. The bR double cysteine mutant D38C/ F156C (Strain L133) was kindly provided by Janos Lanyi (UC Irvine). Expression and purification of the mutant followed standard protocols.49 SDSL of bR D38C/F156C in the PM was achieved as previously described with the modification that spin-labeling was performed at RT rather than 4 °C.50 The MTSL-modified bR D38C/F156C (D38R1/F156R1) was brought to a final concentration of 400 μM in the same buffer with the addition of 30% (w/v) glycerol and flash-frozen in liquid nitrogen. DMPC vesicles were formed by hydrating a lipid film in buffer (50 mM Tris, 300 mM NaCl, pH 8) to a final concentration of 30 mM (20 mg/mL) followed by 10× freeze− thaw cycles and extrusion through a 400 nm filter. wt bR or spin-labeled bR (D38R1/F156R1) in the PM (4 mg) was pelleted by centrifugation (30 m; 40 000 g; RT) and resuspended (1 mL) in the DMPC-liposomes at a final protein/lipid molar ratio of 1:172. The solution was sonicated for 30 m in a bath sonicator with gentle heating.13 1 mL of a 2.5% (w/v) SMA−polymer solution in the same buffer (50 mM Tris, 300 mM NaCl, pH 8) was added dropwise over 2−3 m to the bR-DMPC suspension, and the solution was allowed to equilibrate for 1 h. The resulting bR-Lipodisq solution was centrifuged (30 m; 40 000 g; RT) to partially remove any nonsolubilized bR and purified by size exclusion chromatography (SEC) using a Superdex 200 X16/100 column (GE Healthcare). The resulting purple fractions with an absorbance at 555 nm were pooled for further use. The preparation of solubilized bR was carried out as previously described.38,51 bR (WT or D38R1/F156R1) in the PM (10 mg) was pelleted by centrifugation (30 m; 40 000 g; 4 °C) and resuspended in 6 mL of 25 mM NaiHiPO4, pH 6.9 to which 2 mL of 10% (w/v) OG in water was added. The solution was sonicated (1 m) in a bath sonicator and incubated at RT overnight without stirring. The solution was adjusted to pH 5.5 using 0.1 M HCl and centrifuged (45 m; 100 000 g; 15 °C) to remove any nonsolubilized and aggregated material. The sample was concentrated to 2 mL and applied to a preparative gel column (Sephadex G-75, XK 16/100) with 1.2% OG in 25 mM NaiHiPO4, pH 5.5. Purple fractions containing solubilized bR were pooled and concentrated to a final concentration of 9 mg/m using a using a Vivaspin concentrator with a 50 kDa cutoff (Sartorius Stedium). Absorbance spectra were measured on a Jasco V-630 UV at RT. DLS measurements were performed on a Viscotek 801 particle sizer (Malvern Instruments) at 25 °C. Data represent the average of ten 20 s experiments. A scattering intensity distribution was generated from the data using the software OmniSize 3.0. Circular dichroism (CD) measurements were carried out on a Jasco J815 spectropolarimeter at 18 °C. A 5 mm path length quartz cuvette was used for bR in the PM and in Lipodisq nanoparticles, and a 2 mm path length quartz cuvette was used for bR incorporated into DMPC vesicles. Spectra were recorded between 350 and 700 nm. The digital

■

ASSOCIATED CONTENT

S Supporting Information *

cw EPR distance measurements results are given. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Present Addresses ∥

Department of Biochemistry and Biophysics, Stockholm University, SE-106 91 Stockholm, Sweden. ⊥ Department of Chemistry, University of Birmingham, Edgbaston Birmingham B15 2TT, United Kingdom. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We acknowledge Dr. Jeffrey Harmer and Peter Fisher for technical assistance, Malvern Cosmeceutics for supplying the SMA polymer, and Prof. Janos Lanyi for providing double bR mutant used in this study. Studentship funding was provided by the EPRSC, the Overseas Research council, and the University of Oxford Biochemistry Department (MRO). J.E.L. thanks the Royal Society for a URF. EPSRC are thanked for funding the Centre for Advanced Electron Spin Resonance (EP/D048559/ 1).

■

REFERENCES

(1) Tate, C. G. Methods Mol. Biol. 2010, 601, 187−203. (2) Prive, G. G. Methods 2007, 41, 388−397. (3) Sachs, J. N.; Engelman, D. M. Annu. Rev. Biochem. 2006, 75, 707− 712. (4) Seddon, A. M.; Curnow, P.; Booth, P. J. Biochim. Biophys. Acta 2004, 1666, 105−117. (5) De Angelis, A. A.; Nevzorov, A. A.; Park, S. H.; Howell, S. C.; Mrse, A. A.; Opella, S. J. J. Am. Chem. Soc. 2004, 126, 15340−15341. (6) De Angelis, A. A.; Opella, S. J. Nat. Protoc. 2007, 2, 2332−2338. (7) Bayburt, T. H.; Sligar, S. G. Protein Sci. 2003, 12, 2476−2481.

4691

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692

Nano Letters

Letter

(8) Raschle, T.; Hiller, S.; Etzkorn, M.; Wagner, G. Curr. Opin. Struct. Biol. 2010, 20, 471−479. (9) Geertsma, E. R.; Nik Mahmood, N. A.; Schuurman-Wolters, G. K.; Poolman, B. Nat. Protoc. 2008, 3, 256−266. (10) Fang, G.; Friesen, R.; Lanfermeijer, F.; Hagting, A.; Poolman, B.; Konings, W. N. Mol. Membr. Biol. 1999, 16, 297−304. (11) Vold, R. R.; Prosser, R. S. J. Magn. Reson., Ser. B 1996, 113, 267−271. (12) Bayburt, T. H.; Sligar, S. G. FEBS Lett. 2010, 584, 1721−1727. (13) Knowles, T. J.; Finka, R.; Smith, C.; Lin, Y. P.; Dafforn, T.; Overduin, M. J. Am. Chem. Soc. 2009, 131, 7484−7485. (14) Tonge, S. R. Malvern Cosmeceutics Ltd. Patent GB 2426703, 2007, WO 2006129127, 2006. (15) Jamshad, M.; Lin, Y. P.; Knowles, T. J.; Parslow, R. A.; Harris, C.; Wheatley, M.; Poyner, D. R.; Bill, R. M.; Thomas, O. R.; Overduin, M.; Dafforn, T. R. Biochem. Soc. Trans. 2011, 39, 813−818. (16) Orwick, M.; Judge, P.; Procek, J.; Lindholm, L.; Graziadei, A.; Engel, A.; Gröbner, G.; Watts, A. Angew. Chem., Int. Ed. 2012, DOI: 10.1002/anie.201201355. (17) Reyenolds, J. A.; Stoeckenius, W. Proc. Natl. Acad. Sci. U.S.A. 1977, 74, 2803−2804. (18) Bayburt, T. H.; Grinkova, Y. V.; Sligar, S. G. Arch. Biochem. Biophys. 2006, 450, 215−222. (19) Heyn, M. P.; Bauer, P. J.; Dencher, N. A. Biochem. Biophys. Res. Commun. 1975, 67, 897−903. (20) Cassim, J. Y. Biophys. J. 1992, 63, 1432−1442. (21) Cherry, R. J.; Muller, U. J. Mol. Biol. 1978, 121, 283−298. (22) Lentz, B. R.; Freire, E.; Biltonen, R. L. Biochemistry 1978, 17, 4475−4480. (23) Chen, S. C.; Sturtevant, J. M. Biochemistry 1981, 20, 713−718. (24) Sternberg, B.; L’Hostis, C.; Whiteway, C. A.; Watts, A. Biochim. Biophys. Acta 1992, 1108, 21−30. (25) Sternberg, B.; Watts, A.; Cejka, Z. J. Struct. Biol. 1993, 110, 196−204. (26) Negishi, L.; Mitaku, S. J. Biochem. 2011, 150, 113−119. (27) Berliner, L. J.; Grunwald, J.; Hankovszky, H. O.; Hideg, K. Anal. Biochem. 1982, 119, 450−455. (28) Kusnetzow, A. K.; Altenbach, C.; Hubbell, W. L. Biochemistry 2006, 45, 5538−5550. (29) Mitsuoka, K.; Hirai, T.; Murata, K.; Miyazawa, A.; Kidera, A.; Kimura, Y.; Fujiyoshi, Y. J. Mol. Biol. 1999, 286, 861−882. (30) Grigorieff, N.; Ceska, T. A.; Downing, K. H.; Baldwin, J. M.; Henderson, R. J. Mol. Biol. 1996, 259, 393−421. (31) Pfeiffer, M.; Rink, T.; Gerwert, K.; Oesterhelt, D.; Steinhoff, H. J. J. Mol. Biol. 1999, 287, 163−171. (32) Rink, T.; Riesle, J.; Oesterhelt, D.; Gerwert, K.; Steinhoff, H. J. Biophys. J. 1997, 73, 983−993. (33) Lanyi, J. K. Annu. Rev. Physiol. 2004, 66, 665−688. (34) Goldman, S. A.; Freed, J. H.; Bruno, G. V. J. Phys. Chem. 1972, 76, 1858. (35) Freed, J. H. In Spin Labeling. Theory and Applications, Vol. 1; Berliner, L. J., Ed.; Academic Press: New York, 1976; pp 53−132. (36) Hubbell, W. L.; McHaourab, H. S.; Altenbach, C.; Lietzow, M. A. Structure 1996, 4, 779−783. (37) McHaourab, H. S.; Lietzow, M. A.; Hideg, K.; Hubbell, W. L. Biochemistry 1996, 35, 7692−7704. (38) Nollert, P. Methods 2004, 34, 348−353. (39) Reginsson, G. W.; Schiemann, O. Biochem. J. 2011, 434, 353− 363. (40) Jeschke, G.; Polyhach, Y. Phys. Chem. Chem. Phys. 2007, 9, 1895−1910. (41) McHaourab, H. S.; Steed, P. R.; Kazmier, K. Structure 2011, 19, 1549−1561. (42) Endeward, B.; Butterwick, J. A.; MacKinnon, R.; Prisner, T. F. J. Am. Chem. Soc. 2009, 131 (42), 15246−15250. (43) Banham, J. E.; Baker, C. M.; Ceola, S.; Day, I. J.; Grant, G. H.; Groenen, E. J. J.; Rodgers, C. T.; Jeschke, G.; Timmel, C. R. J. Magn. Reson. 2008, 191, 202−218.

(44) Steinhoff, H. J.; Radzwill, N.; Thevis, W.; Lenz, V.; Brandenburg, D.; Antson, A.; Dodson, G.; Wollmer, A. Biophys. J. 1997, 73, 3287−3298. (45) Steinhoff, H. J. Front Biosci. 2002, 7, C97−C110. (46) Polyhach, Y.; Jeschke, G. Spectroscopy 2010, 24, 651−659. (47) Zou, P.; McHaourab, H. S. Biophys. J. 2010, 98, L18−20. (48) Hirai, T.; Subramaniam., S. FEBS Lett. 2003, 545, 2−8. (49) Oesterhelt, D.; Stoeckenius, W. Methods Enzymol. 1974, 31, 667−678. (50) Eliash, T.; Weiner, L.; Ottolenghi, M.; Sheves, M. Biophys. J. 2001, 81, 1155−1162. (51) Landau, E. M.; Rosenbusch, J. P. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 14532−14535. (52) Jeschke, G. ChemPhysChem 2002, 3, 927−932. (53) Jeschke, G.; Chechik, V.; Ionita, P.; Godt, A.; Zimmermann, H.; Banham, J.; Timmel, C. R.; Hilger, D.; Jung, H. Appl. Magn. Reson. 2006, 30, 473−498.

4692

dx.doi.org/10.1021/nl3020395 | Nano Lett. 2012, 12, 4687−4692