Crystallogenesis of Membrane Proteins Mediated by Polymer-Bounded Lipid Nanodiscs

Transcription

1 Resource Crystallogenesis of Membrane Proteins Mediated by Polymer-Bounded Lipid Nanodiscs Graphical Abstract Authors Jana Broecker, Bryan T. Eger, Oliver P. Ernst Correspondence (J.B.), (O.P.E.) In Brief Broecker et al. describe extraction and purification of a membrane protein embedded in a polymer-bounded lipid nanodisc and its transfer into a lipid environment suited to grow welldiffracting crystals. Without ever exposing the protein to conventional detergents, a high-quality, highresolution membrane-protein structure can be obtained. Highlights d Crystallization of a membrane protein that never left a lipid bilayer environment Accession Numbers 5ITE 5ITC d d d Lipid nanodiscs enable transfer of membrane proteins between lipid bilayer systems SMA copolymers are compatible with lipidic cubic phases Method expands the toolbox for membrane-protein crystallography Broecker et al., 2017, Structure 25, February 7, 2017 ª 2017 Elsevier Ltd.

2 Structure Resource Crystallogenesis of Membrane Proteins Mediated by Polymer-Bounded Lipid Nanodiscs Jana Broecker, 1, * Bryan T. Eger, 1 and Oliver P. Ernst 1,2,3, * 1 Structural Neurobiology, Department of Biochemistry 2 Department of Molecular Genetics University of Toronto, 1 King s College Circle, Toronto, ON M5S 1A8, Canada 3 Lead Contact *Correspondence: jana.broecker@utoronto.ca (J.B.), oliver.ernst@utoronto.ca (O.P.E.) SUMMARY For some membrane proteins, detergent-mediated solubilization compromises protein stability and functionality, often impairing biophysical and structural analyses. Hence, membrane-protein structure determination is a continuing bottleneck in the field of protein crystallography. Here, as an alternative to approaches mediated by conventional detergents, we report the crystallogenesis of a recombinantly produced membrane protein that never left a lipid bilayer environment. We used styrene maleic acid (SMA) copolymers to solubilize lipid-embedded proteins into SMA nanodiscs, purified these discs by affinity and size-exclusion chromatography, and transferred proteins into the lipidic cubic phase (LCP) for in meso crystallization. The 2.0-Å structure of an a-helical seven-transmembrane microbial rhodopsin thus obtained is of high quality and virtually identical to the 2.2-Å structure obtained from traditional detergent-based purification and subsequent LCP crystallization. INTRODUCTION Biological membranes contain a great variety of membrane proteins, which fulfill vital functions as receptors, signal transducers, channels, transporters, motors, and anchors (Fiedler et al., 2010). High-resolution structures of membrane proteins are still rare but are of paramount importance for understanding protein function on a molecular level with significant implications for biomedical and pharmaceutical applications (Bill et al., 2011). For in vitro studies such as biophysical analyses and structure determination, membrane proteins are usually purified with the aid of detergents. However, micelles formed by conventional detergents are poor membrane mimetics (Helenius and Simons, 1975), and membrane proteins in micelles do not experience the same lateral pressure profile as in a lipid bilayer (Van den Brink-van der Laan et al., 2004), which in many cases is important to maintain the native fold and function of the protein. Moreover, detergents tend to remove annular lipids, which are often essential for membrane-protein function (Paila et al., 2009). Accordingly, many membrane proteins are inactive in detergent micelles and require a membrane environment in order to be functional. Moreover, structural heterogeneity and conformational instability of detergent-solubilized membrane proteins can complicate quantitative as well as structural analyses (Privé, 2007). As alternatives, several peptide-based amphiphiles (e.g., peptitergents, Schafmeister et al., 1993; lipopeptide detergents, McGregor et al., 2003; or short peptide surfactants, Zhao et al., 2006) as well as amphipols (Tribet et al., 1996) and fluorinated surfactants (Abla et al., 2015; Frotscher et al., 2015) have been explored, but so far have not gained broad acceptance. In the last years, methods based on fragmenting membranes into nanometer-scale particles have revolutionized membrane-protein research (Rajesh et al., 2011). Depending on how such particles are reconstituted, bicelles (D urr et al., 2012), membrane scaffold protein (MSP) nanodisc (Bayburth and Sligar, 2003), and saposin lipoprotein (Salipro) nanoparticles (Frauenfeld et al., 2016) can be distinguished. However, neither of these approaches is designed to extract proteins from biological membranes. Thus, they require detergent-solubilized protein and are still impaired by the limitations described above. Recently, polymer-bounded lipid nanodiscs have been introduced as a novel, native-like, nanometer-sized membrane system that opens new perspectives for the in vitro characterization of membrane proteins (Knowles et al., 2009; for a review see Dörr et al., 2016). SMA copolymers, which comprise styrene (S) and maleic acid (MA) building blocks, self-assemble with lipids into nanoparticles (Vargas et al., 2015) and solubilize membrane proteins from cellular membranes without the need of conventional, head-and-tail detergents. While conventional detergents solubilize proteins and lipids into micelles, SMA copolymers wrap around planar lipid bilayer fragments containing membrane proteins and their surrounding lipids, forming so-called SMA nanodiscs, which, in some cases, have been shown to better preserve membrane-protein function and thus fold (Knowles et al., 2009; Dörr et al., 2014; Gulati et al., 2014; Swainsbury et al., 2014). SMA nanodiscs contain mainly lipids of the protein s microenvironment (Dörr et al., 2014) and, owing to their small size, are amenable to various biophysical techniques (Lee et al., 2016). Here, we report the crystallogenesis of a membrane protein that never left a lipid bilayer since its production in the cell. We used SMA nanodiscs as vehicles for the fast, cheap, and mild transfer of a recombinantly produced membrane protein from its cellular membrane environment into the lipidic cubic phase 384 Structure 25, , February 7, 2017 ª 2017 Elsevier Ltd.

3 Figure 1. Detergent-Based versus Polymer- Mediated in Meso Crystallogenesis Comparison of membrane-protein crystallization in the LCP following (left/blue) traditional detergentbased protein solubilization and purification or (right/yellow) polymer-based protein solubilization and purification retaining a bilayer environment. As proof of principle, a seven-transmembrane a-helical membrane protein is produced in E. coli and either solubilized and purified with detergent (blue; step 1) or solubilized and purified with SMA copolymers (yellow; step 1). Solubilized proteins are purified chromatographically (step 2) and transferred into the LCP (step 3) for growth of crystals (step 4). Protein structures are solved by X-ray crystallography (step 5). Lipid molecules are shown in green. SMA-LCP approach is of high quality and virtually identical to a 2.2-Å structure obtained from a traditional detergent LCP approach. RESULTS (LCP), from which well-diffracting crystals can be grown (Figure 1). The LCP (or in meso) method has emerged as one of the most popular approaches for growing membrane-protein crystals, boosting the number and resolution of membrane-protein structures (Landau and Rosenbusch, 1996; Caffrey and Cherezov, 2009). Commonly, the monoacylglycerol monoolein forms the curved bicontinuous LCP bilayer, from which proteins in the presence of precipitants preferentially partition into lamellar domains and form crystals (Caffrey, 2000; Caffrey et al., 2012). We exemplified the polymer-mediated crystallization technology by solving the crystal structure of the proton pump bacteriorhodopsin from Haloquadratum walsbyi (HwBR) (Hsu et al., 2015), which can be recombinantly produced in Escherichia coli in quantities sufficient for crystallization. Rhodopsins are seven-transmembrane a-helical proteins with a covalently bound retinal chromophore, which renders the proteins photochemically active (Ernst et al., 2014). For direct comparison, we devised two crystallization strategies (Figure 1), starting with protein solubilization and purification using either n-dodecyl-b-d-maltoside (DDM) and n-octyl-b-d-glucoside (OG) as traditional detergents or SMA copolymers, which yielded HwBR preparations that were never depleted of a lipid bilayer environment. His-tagged HwBR was purified by immobilizedmetal affinity chromatography (IMAC) followed by size-exclusion chromatography (SEC) and transferred into the LCP for crystallization. From well-ordered crystals, high-resolution protein structures were determined by means of X-ray crystallography under cryogenic conditions. The 2.0-Å structure thus obtained from the Membrane-Protein Solubilization and Purification To solubilize HwBR proteins produced in E. coli, we broke the cells and isolated E. coli inner cell membranes by differential centrifugation. As is often the case for traditional detergent-based methods, we had to employ two different detergents for HwBR: DDM micelles for solubilizing and OG micelles for purifying and crystallizing the reference sample. When solubilizing the membranes with SMA copolymers, we found that the protein yield for HwBR was low. To optimize solubilization efficiency, we modulated bilayer characteristics such as bilayer composition, temperature, ph, ionic strength, and co-solutes (Note S1 and Figures S1A S1B) and found that the addition of externally added synthetic phospholipid 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) increased the yield of solubilized HwBR to an overall yield close to that of DDM solubilization. The terms SMA approach or polymer-mediated approach are used henceforth to indicate the use of SMA and DMPC for solubilization. Protein-containing SMA nanodiscs have been purified to substantial degrees by IMAC using affinity tags attached to the membrane proteins and subsequent SEC (Dörr et al., 2014; Gulati et al., 2014; Jamshad et al., 2015). However, as indicated previously (Dörr et al., 2014; Gulati et al., 2014), hexahistidine tags, His 6 -tags, did not properly allow us to bind HwBR-containing SMA nanodiscs to Ni 2+ and Co 2+ resins. Thus, we extended the standard His 6 -tag to a double His 6 -tag (see Supplemental Experimental Procedures) for the efficient separation of HwBR embedded in SMA nanodiscs from SMA nanodiscs that did not contain the target proteins by Ni 2+ IMAC. As a second purification step we performed SEC (Figure 2B). Purity of proteins purified in either OG detergent micelles or in SMA nanodiscs was comparable (Figure S1C). For the SMA approach, this Structure 25, , February 7,

4 Figure 2. Biophysical Characterization of HwBR in OG Micelles or SMA Nanodiscs Results of solubilization optimization with SMA are given in Figures S1A and S1B. (A) SDS PAGEs reflecting the purity of both preparations after Ni 2+ -based IMAC and SEC with the HwBR band (29.3 kda) and some impurities (*,**) indicated (M, marker; MP, main peak; SP, side peak). For a preparation with higher purity see Figure S1C. (B) SEC profiles for HwBR in OG micelles versus SMA nanodiscs (A, absorbance; V, volume). The latter are larger and less homogeneous than HwBR OG complexes. (C) UV-visible absorbance spectra of purified HwBR. SMA copolymers contribute significantly to the A 280nm signal. Absorbance at 420 nm is likely due to co-purified soluble hemeproteins (l, wavelength). (D) Intensity-weighted size distributions obtained from dynamic light scattering (I, intensity; d, hydrodynamic diameter). required an additional Co 2+ IMAC that reduced the protein yield. To overcome the lower protein yield and because LCP crystallization can tolerate the presence of impurities to some degree (Kors et al., 2009), we preferred working with a less-pure protein sample (Figure 2A) that, in comparison with the purer sample, gave similar crystallization hits. Biophysical Characterization of Proteins in Detergent Micelles and SMA Nanodiscs Proteins in both preparations, purified in either OG detergent micelles or in SMA nanodiscs, showed an absorbance maximum at 550 nm typical of covalently bound all-trans-retinal in the chromophore binding pocket of HwBR (Figure 2C). The apparent hydrodynamic diameter of HwBR-containing SMA nanodiscs was about 18 nm, as derived from dynamic light scattering (DLS; Figure 2D). In previous studies, DLS-based SMA nanodisc sizes of about 12 nm have been reported (Knowles et al., 2009; Orwick- Rydmark et al., 2012; Swainsbury et al., 2014), which could indicate that in our preparation the majority of nanodiscs are actually smaller than 18 nm and that some of them might be stacked or associated. Membrane-Protein Crystallization from Detergent Micelles and SMA Nanodiscs HwBR proteins were transferred from OG micelles or SMA nanodiscs, respectively, and reconstituted into the LCP by homogenizing HwBR OG micelles or protein-loaded SMA nanodiscs, respectively, with an excess of monoolein (Caffrey and Cherezov, 2009). The protein-to-lipid ratio was 2:3 (v/v) and, thus, lipid hydration was 60% (see Experimental Procedures and Table S1). Crystallization conditions in the LCP were optimized individually for HwBR transferred from either OG micelles or SMA nanodiscs (Table S1). In both cases, we first screened several hundred crystallization conditions using various commercially available screens. We then systematically optimized several hit conditions before performing a second screening, for which we added individual chemicals to the best-hit conditions (Experimental Procedures). The number of crystallization hits obtained for both approaches was comparable, and when SMA was present the LCP bolus remained non-birefringent when viewed between crossed polarizers. Also, we noted an increase in the viscosity of the forming LCP when mixing monoolein and HwBR in SMA nanodiscs. This affected the distribution of the LCP on to glass plates using an LCP robot and required marginal changes to the protocol (Experimental Procedures). In addition, precautions had to be taken when harvesting crystals, because the LCP was even more viscous than usual. Note that several in situ data collection technologies exist that eliminate the need for harvesting membrane-protein crystals from the viscous LCP (Huang et al., 2015, 2016; Broecker et al., 2016). In the LCP, both OG- and SMA-purified HwBR preparations yielded similarly shaped, flat crystals (Figure S2 and Table S1) with identical unit cell dimensions (Table S2), which diffracted to 2.0 Å (detergent LCP approach; data not shown) and 1.8 Å (SMA LCP approach) resolution, respectively (Figure 3). Most importantly, the presence of SMA copolymers did not adversely affect crystallization parameters such as crystal packing or shape, lipid-phase behavior, or diffraction quality (Table S1). Conventional Detergent and SMA Yield Virtually Identical HwBR Structures We selected individual best crystals, collected data at the Advanced Photon Source synchrotron, and determined the atomic structures by molecular replacement followed by refinement of the atomic models. Overall, HwBR structures obtained from the detergent-based and polymer-mediated approaches are virtually identical (Table S2), and also agree well with a recently solved structure obtained from HwBR crystals grown in the LCP after purification in n-decyl-b-d-maltoside (DM) (Hsu et al., 2015). In all cases, the asymmetric unit cell contains a trimer (Figure S3A), and monoolein molecules at the monomer interfaces mediate trimer formation (Figure S3B). For the two structures determined here, the root-mean-square deviation (RMSD) of C a atoms is 0.22 Å (Figure S3F). Larger RMSD values correlate well with higher crystallographic B factors (Figure S3G), the result of weaker electron density observed for the termini and flexible loop regions of the proteins. In the living cell, HwBR pumps protons from the cytoplasm to the cell exterior, ultimately resulting in the production of ATP by ATP synthase (Figure 4A). Key residues that are part of the proton translocation path are placed in identical positions in both 386 Structure 25, , February 7, 2017

5 Figure 3. Diffraction Quality of Crystals Generated by the Polymer- Mediated Approach Crystals obtained from protein purified in SMA nanodiscs and grown in the LCP diffract to a resolution better than 2.0 Å. (A) Typical X-ray diffraction pattern as detected using a PILATUS detector at the Advanced Photon Source at Argonne National Laboratory (Lemont, IL, USA). Note that a pattern with one well-defined and one diffuse ring is characteristic of crystals grown in the LCP. Three typical intense, round spots are shown at the right (frame colors match those in the complete pattern). (B) Four examples of reflection spots (arrows) with a resolution better than 2.0 Å that can still be seen by the naked eye. Any standard indexing software is even more sensitive than visual inspection and will detect more spots at resolutions better than those indicated. Crystal images are shown in Figure S2. structures (Figures 4B 4D). Residues in the retinal-binding pockets superimpose, and all-trans-retinals are covalently bound to residues Lys224 (Figure 4E), as expected for the HwBR ground state. The retinal ligands are well resolved, and their omission during refinement generates clear positive density in the resulting F o F c difference maps (data not shown). Taken together, an integrated approach using SMA nanodiscs and crystallization in the LCP enabled us to obtain the atomic structure of the seven-transmembrane a-helical protein HwBR in a lipidic environment under native-like conditions. The SMA copolymer does not compromise diffraction quality and electron density maps. Indeed, the values of key quality indicators of the corresponding structure confirm that a structure of high quality can be obtained by the SMA LCP approach (PDB: 5ITC). DISCUSSION Despite many advances in the field, determining the structure of membrane proteins by X-ray crystallography is still a major challenge; as a consequence, the number of high-resolution membrane-protein structures still lags far behind that of soluble proteins. New detergents that better stabilize membrane proteins Figure 4. Structural Alignment of HwBR Monomers Structural alignment of HwBR monomers obtained from LCP-grown crystals that originated either from protein solubilized in DDM micelles and purified in OG micelles (green; all-trans-retinal shown in blue) or solubilized and purified in SMA nanodiscs (magenta; all-trans-retinal shown in orange) with a C a RMSD of 0.22 Å for the trimer in the asymmetric unit. (A D) HwBR pumps protons from the intracellular side (intra) to the extracellular side (extra) (A). Parts of the proton translocation path with functional residues in stick representation are indicated as follows: (B) retinal-binding pocket with all-trans-retinal bound to Lys224 by a Schiff base linkage and with proton re-uptake residue Asp93, (C) proton-releasing complex (note that the Glu202 side chain shows some flexibility in orientation between different protomers), and (D) proton outward cap region with hydrogen bonds shown as gray dashes. (E) The 2F o F c electron density maps of retinal and the surrounding residues are contoured at 1.0s and shown in blue (detergent LCP) and orange (SMA LCP). Comparison and structural alignment of structures obtained from the detergent-based or polymer-mediated approaches are given in Figure S3. Crystallization and crystal information as well as data collection and refinement statistics are given in Tables S1 and S2. have been developed recently (Chae et al., 2010; Ghosh et al., 2015), but screening for the best detergent remains a tedious task, because each membrane protein behaves differently. In addition, and as was the case here, often different detergents are required for the solubilization, stabilization, and/or crystallization of a membrane protein (see Experimental Procedures) (Privé, 2007). Hence, the use of SMA for extracting membrane proteins from cellular membranes of native or heterologous expression hosts and for stabilizing them in a lipid bilayer environment can obviate time-consuming detergent-screening procedures and accelerate the crystallization process in its early stages. Moreover, SMA has to be supplied only for initial solubilization, while all subsequent steps are carried out without addition of more SMA, making the approach described here simple and substantially cheaper than comparable detergentrelated work. Structure 25, , February 7,

6 As is generally the case in protein crystallization, also SMAsolubilized membrane proteins require optimization of LCP crystallization conditions according to standard protocols known from detergent-based work. While the number of crystallization hits, LCP stability, or crystallization parameters, such as crystal packing or shape, were not adversely affected by the presence of SMA, it is difficult to judge whether and how SMA copolymers influence the crystallization process itself. The optimized crystallization conditions differ between the OG LCP and SMA LCP approaches (Table S1), and a third LCP crystallization study on HwBR purified in DM yielded yet another, chemically different crystallization condition (Hsu et al., 2015). This is not surprising, since crystallization in LCP is favored in the presence of reagents that, while also triggering the formation of local lamellar phases, affect the protein s solubility/stability and promote nucleation (Caffrey, 2015). Consequently, crystallization conditions are very sensitive to the environment of the membrane protein. In our experience and when compared with the effect of OG, the presence of SMA copolymers neither negatively nor positively influences the crystallization process. As long as the amount of SMA (or detergent) is kept low, the LCP appears to tolerate the presence of these amphiphiles. Given that SMA copolymers extract a membrane protein with bound lipids, the question arises as to why there were no phospholipids of the expression host revealed in the HwBR structure? The co-purification of lipids with membrane proteins in a traditional detergent-based approach is remarkably different from the co-purification of a larger number of lipids of the microenvironment of the protein in a polymer-bounded nanodisc. Thus, in an SMA nanodisc the lipid environment a protein finds in its expression host is retained, but this does not necessarily mean that these lipids are bound tightly. By contrast, upon detergentmediated membrane-protein extraction, which is typically a harsh process, it is mainly those lipids that are tightly bound to the protein that are co-purified (Valiyaveetil et al., 2002; Shinzawa-Itoh et al., 2007). Thus, the fact that no phospholipids were revealed in both structures indicates that there are no lipids in the proximity of HwBR that interact avidly with the protein. Note that the lipid bilayer composition of Haloquadratum walsbyi, a halophilic microorganism, differs drastically from that of E. coli. For instance, phosphoethanolamine (PE) lipids, which are abundant in E. coli membranes (Furse and Scott, 2016), are not found at all among the total polar lipids extracted from the membranes of H. walsbyi cells (Lobasso et al., 2008). It has been shown that the monoolein cubic phase can accommodate up to 20 mol% of other lipids, provided that monoolein is fully miscible with each of these lipids (Cherezov et al., 2002). If, in our approach, protein is transferred from SMA nanodiscs into the monooleinbased LCP for crystallization, the nanodisc-embedded lipids are transferred as well, and eventually mix with an excess of monoolein lipids. As a consequence, any lipid that is not tightly bound or not necessary for crystal packing might be substituted with a monoolein lipid. In the particular case of HwBR, we noticed that monoolein has a tendency to fill empty or open spaces in the protein structure, thereby supporting the assembly of monomers into trimers and also assisting in crystal packing (Figure S3B). The addition of DMPC to aid the solubilization process of a-helical membrane proteins is a modification to existing protocols and has been suggested previously (Dörr et al., 2016). Notably, a protein-rich membrane can be difficult to solubilize. Thus, externally added DMPC might help in the solubilization process simply because the polymer needs a certain amount of lipids for solubilization and formation of nanodiscs. Other specific factors could be involved as well. For instance, DMPC lipids might modulate the lateral pressure profile in the E. coli membrane (Fiedler et al., 2010), which in contrast to cell membranes of other expression hosts, such as yeast or mammalian cells, where DMPC addition during solubilization appears not to be necessary (Long et al., 2013; Jamshad et al., 2015), contains up to 80% of PE lipids (Furse and Scott, 2016). The presence of PE lipids with small head groups in a lipid bilayer redistributes positive lateral pressure from the head group region to the hydrocarbon core, slowing down SMA-mediated membrane solubilization (Scheidelaar et al., 2015). Addition of cylindrically shaped DMPC lipids might counteract this effect. Note that while the successful solubilization of a membrane protein from the outer membrane of E. coli using SMA has not been reported so far, PagP could be solubilized from DMPC proteoliposomes (Knowles et al., 2009), indicating that DMPC might also aid the solubilization process in the case of b-barrel membrane proteins produced in E. coli. For the transfer of membrane proteins from SMA nanodiscs into the LCP, we propose the following working model: SMA copolymer chains, which initially wrap around nanodiscs, partition into the monooleins. As a consequence, nanodisc-embedded lipids and membrane proteins are transferred into the monoolein membrane. Various studies have shown that SMA copolymers non-preferentially extract different phospholipid species from native (Long et al., 2013; Swainsbury et al., 2014; Prabudiansyah et al., 2015) or model membranes (Scheidelaar et al., 2015; Cuevas et al., 2016). While we do not know whether SMA copolymers prefer monooleins to phospholipids (the partition coefficients of SMAs for monoolein remain to be determined), preferences for a specific lipid are likely negligible when monoolein is present in excess. Accordingly, phospholipid molecules exchange between SMA nanodiscs and vesicular model membranes (Cuevas et al., 2016) or lipid monolayers (Hazell et al., 2016) by addition of excess lipid. Thus, we assume that transfer of lipids and HwBR proteins from SMA nanodiscs into the LCP is triggered by the addition of the excess of monoolein that is typically used in an LCP experiment. Based on this model, there are two possible explanations why SMA HwBR-loaded LCP showed increased viscosity over OG HwBR-loaded LCP. The absence of elevated concentrations of detergents may increase viscosity. However, we did not experience this when setting up crystallization trials with water-soluble proteins in the absence of detergents, rendering this explanation unlikely. Alternatively, when SMA copolymers partition into the excess monoolein phase, they bind to the membrane surface, where they can interact with a multitude of lipid head groups and thus may change the lipid-phase behavior in favor of a more gel-like phase. Halobacterium salinarum bacteriorhodopsin has been incorporated before into the LCP from native purple membranes in a completely detergent-free manner (Nollert et al., 1999). However, this is a unique case, because the vast majority of membrane proteins are naturally not found in high densities as in purple membranes but, instead, require recombinant protein production and chromatographic purification in a detergentsolubilized state before crystallization. In contrast to this very 388 Structure 25, , February 7, 2017

7 specific case, we demonstrate a more general, polymer-mediated crystallization technology, in which a recombinantly produced membrane protein can be extracted and chromatographically purified in SMA-bounded lipid nanodiscs to render it amenable to in meso crystallization without the need of conventional detergents at any time. The question arises whether protein-containing SMA-bounded or MSP-bounded lipid nanodiscs can be crystallized directly. In our hands and to our knowledge, such nanodiscs have never been shown to give rise to well-diffracting membrane-protein crystals. On a more general note, while in one report a single copy of a channel protein has been reconstituted from an SMA nanodisc into a planar lipid bilayer (Dörr et al., 2014), we demonstrate the mass transfer of membrane proteins from SMA nanodiscs into another lipid bilayer system, which will also be of interest for those studying proteins in, for instance, lipid vesicles. SMA-based extraction is applicable to a wide variety of membrane proteins, ranging from single-membrane-spanning a-helices (Paulin et al., 2014) to oligomeric membrane-protein complexes with up to 36 transmembrane helices (Postis et al., 2015). The broad applicability of SMA copolymers for the isolation of membrane proteins together with their efficient transfer into an LCP environment for crystal growth, as presented here, bears the potential for more structures of membrane proteins to become available, which so far have not been amenable to solubilization in detergents. The bilayer environment is retained in SMA nanodiscs, even though their lipid core exerts an altered lateral pressure profile with respect to the situation in a vesicular bilayer membrane (Orwick et al., 2012). However, being embedded in a membrane-like environment might be responsible for the fact that in several cases SMA nanodiscs have proven to be the tool of choice for stabilizing membrane proteins (Dörr et al., 2014, 2016; Swainsbury et al., 2014). Recently, also notoriously challenging G-protein-coupled receptors have been successfully isolated in their functional forms with the aid of SMA copolymers (Jamshad et al., 2015; Logez et al., 2016). In the case of human adenosine A 2A receptor, SMA nanodiscs increased the thermostability of the receptor by 5.5 C compared with a detergent micelle (Jamshad et al., 2015). Thus, we are optimistic that for some membrane proteins, which for crystallization would require thermostabilizing mutations or engineered disulfide bridges, SMA nanodiscs might reduce the amount of these sequence modifications. Note that, by contrast, the method presented here will not reduce the need for sequence modifications that are important for the crystallization process itself, this is modifications reducing conformational flexibility and/or stabilizing a particular conformation of the membrane protein (Ghosh et al., 2015). Our approach together with low-temperature LCP methods (Li et al., 2013; Salvati Manni et al., 2015), in particular, may thus pave the way to more pristine structures of membrane proteins with less sequence manipulation. Since HwBR is a rather stable protein, the question remains whether the strategy presented here will be applicable to more challenging membrane proteins. On the one hand, SMA copolymers bring along the advantages mentioned above. On the other hand, it is clear that SMA copolymers have peculiarities, not all of which are favorable for membrane-protein crystallogenesis (for a review, see Dörr et al., 2016). For instance, they absorb in the UV range and hamper A 280 nm measurements. However, reliable estimates of protein concentration can be obtained for proteins purified in SMA nanodiscs (Lee et al., 2016). Also, as polyacids, SMA copolymers have metal-chelating properties, resulting in a limited solubility in the presence of multivalent cations. Finally, SMA copolymers demonstrate a restricted ph range over which they function as solubilizing agents. However, improved MAbased copolymers will be introduced shortly that display a low extinction coefficient at 280 nm, have reduced metal-chelating properties, and work over an extended ph range. Taken together, SMA copolymers in combination with LCP add a new and powerful tool to the crystallographer s toolbox, and it is conceivable that the methodology presented here, particularly because of its simplicity, might be used for crystallization of other membrane proteins in a similar way using SMA or other nanodisc-forming polymers. EXPERIMENTAL PROCEDURES If not stated otherwise, all methods were done at room temperature. Materials and SMA Copolymers For details on materials see Supplemental Experimental Procedures. SMA copolymers with a molar styrene-to-maleic acid ratio of 3:1 (mass-average molar mass of M w = 10 kg/mol, number-average molar mass of M n = 4 kg/mol) was obtained as sodium salt solution (XIRAN SL25010 S25) from Polyscope. SMA stock solutions (6%, w/v) were prepared in unadjusted 50 mm Tris buffer, and the ph was adjusted to neutrality by gradually adding HCl. Stock solutions were stored at 20 C and adjusted to ph 8.0 after thawing. UV-Visible Spectroscopy and DLS Measurements For details see Supplemental Experimental Procedures. Generation of Plasmids and Protein Production For details see Supplemental Experimental Procedures. Protein Solubilization from E. coli Membranes Using Detergents For all steps, exposure to light was avoided as much as possible. About 8 g of cells (wet weight) was resuspended in 50 ml of breakage buffer (50 mm 2-(N-morpholino)ethanesulfonic acid [MES] [ph 6.5], 500 mm NaCl) with freshly added EDTA-free protease inhibitors and 1,250 units Benzonase and disrupted by homogenization. The volume was adjusted to 100 ml, and DDM was added to a final concentration of 1.5% (w/v). The suspension was incubated overnight with gentle agitation at 4 C. The next day, solubilized material was isolated by ultracentrifugation at 100,000 3 g and 4 C for 60 min. Protein Solubilization from E. coli Membranes Using SMA Copolymers For all steps, exposure to light was avoided as much as possible. About 8 g of cells (wet weight) was resuspended in 50 ml of breakage buffer (50 mm tris(hydroxymethyl)aminomethane [Tris] [ph 8] and 100 mm NaCl) with freshly added EDTA-free protease inhibitors and 1,250 units of Benzonase and disrupted by homogenization. Cell debris was removed by low-speed centrifugation at 10,000 3 g and 4 C for 30 min. Crude cell membranes were collected by ultracentrifugation at 150,000 3 g and 4 C for 45 min and stored on ice. Membranes were resuspended in solubilization buffer (50 mm Tris [ph 8] and 150 mm NaCl) using a 5-mL disposable syringe and a 25-gauge needle. The final volume was 4.2 ml. For solubilization of membrane proteins, buffer stock solution, NaCl (as powder), and DMPC lipid (as powder) were added. The mixture was sonicated gently (Sonifier S-450; Branson Ultrasonics; settings: output control 3 4, duty cycle 30% 40%, three pulses of 10 s each, samples were cooled between pulses) until the suspension became clear. Alternatively, DMPC liposomes were added. Then SMA copolymers (XIRAN SL25010 S25; as a liquid) were added to give a final solubilization buffer containing 50 mm Tris (ph 8), 300 mm NaCl, 1.5% (w/v) DMPC, and 2.5% (w/v) SMA in a volume of about 7.2 ml. The suspension was incubated overnight Structure 25, , February 7,

8 with gentle agitation. The next day, 20 ml of breakage buffer was added to the mixture, and solubilized material was isolated by ultracentrifugation at 150,000 3 g and 4 C for 20 min. Purification of Detergent-Solubilized Protein For IMAC, solubilized material was supplemented with imidazole (final concentration of 20 mm) and incubated for 2 hr with an equilibrated Ni-NTA Superflow affinity resin (QIAGEN) in the dark. Thereafter, the resin was packed into a column and washed (10 column volumes [CV]) with wash buffer 1 (50 mm MES [ph 6.5], 500 mm NaCl, 0.01% DDM, and 20 mm imidazole) to remove nonspecifically bound proteins. The detergent was exchanged to OG by an extensive wash step (30 CV) with wash buffer 2 (50 mm MES [ph 6.5], 500 mm NaCl, 1% OG, and 20 mm imidazole). Protein detergent complexes were then eluted using elution buffer (50 mm MES [ph 6.5], 500 mm NaCl, 1% OG, and 200 mm imidazole). All fractions containing protein were pooled, concentrated in a 0.5-mL centrifugal filter unit with a molecular weight cutoff (MWCO) of 30 kda, centrifuged (10 C, 5,000 3 g, 10 min), and further purified on a Superdex /300 GL column (GE Healthcare) by SEC using degassed breakage buffer and 1% OG. Fractions of the main peak were collected and concentrated further in centrifugal units with an MWCO of 30 kda to about 200 ml with a concentration of 15 mg/ml as estimated by UV-visible spectroscopy. Protein in OG micelles can be flash-frozen in liquid nitrogen for long-term storage at 80 C. For protein characterization see Supplemental Experimental Procedures. Purification of HwBR SMA Nanodiscs SMA-solubilized material was applied to an equilibrated IMAC column containing Ni-NTA Superflow resin in the dark. Thereafter, the column was washed extensively (10 CV each) with breakage buffer and wash buffer (50 mm Tris [ph 8], 100 mm NaCl, and 20 mm imidazole) to remove non-specifically bound proteins. SMA nanodiscs containing histidine-tagged proteins were then eluted using elution buffer (50 mm Tris [ph 8], 100 mm NaCl, and 500 mm imidazole). All fractions containing protein were pooled, concentrated in a centrifugal filtration unit with an MWCO of 100 kda, centrifuged (10 C, 5,000 3 g, 10 min), and further purified by SEC on a 24-mL Superdex /300 GL column using degassed breakage buffer. Fractions with an A 550nm signal were collected and concentrated further in centrifugal filtration units with an MWCO of 100 kda to about 200 ml with a protein concentration of 13 mg/ml as estimated by UV-visible spectroscopy. Protein in SMA nanodiscs can be flash-frozen in liquid nitrogen for long-term storage at 80 C. For protein characterization see Supplemental Experimental Procedures. LCP Crystallization We followed standardized LCP protocols with minor modifications (Caffrey and Cherezov, 2009). Purified HwBR in either OG detergent micelles (15.5 mg/ml) or SMA nanodiscs (13 mg/ml) was transferred into the LCP with lipid at full lipid hydration (this is, at about 60% hydration). Monoolein was mixed with protein at a 2:3 (v/v) protein-to-lipid ratio. To do so, monoolein was melted at 45 C and then loaded into a 100-mL gas-tight syringe. Protein solution was loaded into another 100-mL syringe, and both syringes were connected through a coupler (Art Robbins Instruments [ARI]). Mesophase was prepared by mixing the contents of both syringes using an LCP mixing station (ARI). With an LCP robot (Gryphon; ARI), protein-loaded mesophase was dispensed on to 96-well glass plates, and precipitant solution was placed on top. While the bolus-to-precipitant ratio was kept constant at 1:5 (v/v), we varied the size of the bolus with a protein/lipid mixture ranging from 0.04 to 0.2 ml. To account for the higher LCP viscosity, lower delivery rates (70%) and smaller gaps between needle and glass plate (0.5 mm) were used when working with the LCP robot. Loaded plates were tightly sealed, wrapped in aluminum foil, incubated at 20 C, and regularly inspected through a stereo microscope (M205C; Leica) equipped with cross-polarizers and an MC170 HD camera (Leica). Crystal-like objects were scored, and protein crystals were distinguished from salt, lipid, or other crystals by comparison with proteinfree crystallization trials and/or UV imaging. Crystallization Conditions Commercially available screens used for an initial screening were: the Cubic Phase I & II Suite (QIAGEN), MembFac HT (Hampton Research), MemGold (Molecular Dimensions), SaltRX HT, PEG/Ion HT, and PEGRx HT (all three Hampton Research). Several initial hits were selected and optimized systematically by slightly changing one condition at a time (e.g., ph value or concentration of buffer, salt, and polyethylene glycols [PEGs]). A secondary screening was performed using following screens (all from Hampton Research): Silver Bullets, Additive Screen, SaltRX HT, PEG/Ion HT, and PEGRx HT. Crystals used for data collection were grown in 8% (v/v) Tacsimate ph 7.0 and 20% (v/v) PEG 3350 in wells with a bolus size of 0.04 ml and 0.2 ml of precipitant solution (HwBR OG) or in 1.6 M NH 4 PO 4 (ph 8.8), 0.1 M NaCl, 0.2 M NaSCN, and 0.016% amino acid mix containing L-His, L-Leu, L-Ile, L-Trp, L-Tyr, and L-Phe in 0.2 M HEPES Na + (ph 6.8) in wells with a bolus size of 0.2 ml and 1.0 ml of precipitant solution (HwBR SMA). Crystals were colored purple and shaped as hexagonal, compact plates or big, flat plates, respectively (Figure S2). The size of the crystals reached about mm 3 within 30 days or mm 3 within 60 days, respectively. Please also refer to Table S1. X-Ray Diffraction Data Collection Crystals were harvested according to Li et al. (2012) and by either using Micro- Grippers with 50 mm aperture (MiTeGen) for HwBR purified in OG micelles or by using Dual-Thickness MicroLoops LD with a 50-mm aperture (MiTeGen) for HwBR purified in SMA nanodiscs. For the SMA LCP approach, crystals were harvested from the extremely viscous LCP particularly carefully so that as little lipid as possible stuck to the loop. Harvested crystals were flash-frozen and stored in liquid nitrogen. X-ray diffraction data were collected at beamlines 17-ID-B and 23-ID-B/D of the Advanced Photon Source(Lemont, IL, USA). A tunable 5 20-mm X-ray beam was used to raster through, locate, and orient the crystal prior to data collection. Crystal parameters were tested using an attenuated beam while optimizing the data collection strategy, and datasets were collected on the best-quality crystals. For details on data collection see Table S2. X-Ray Diffraction Data Analysis Diffraction data of the detergent LCP approach were processed using auto- PROC (Vonrhein et al., 2011), and those of the SMA LCP approach were processed using XDS (Kabsch, 2010) andaimless(evans and Murshudov, 2013). Initial phase information was generated by molecular replacement using the atomic coordinates of HwBR purified in DM detergent micelles as a search model (PDB: 4QI1; Hsu et al., 2015). Before running Phaser (McCoy et al., 2007), bound retinal, monooleins, and waters were removed from the search model. Each dataset yielded a strong solution. Initial refinement was done using Refmac5 (Murshudov et al., 2011) from the CCP4i program package (Winn et al., 2011). In an iterative manner, phases were then improved by structural refinement using Phenix (Adams et al., 2010). The structure was manually examined in Coot (Emsley et al., 2010), and the refined coordinates were rebuilt. Structures were restrained to the reference model (PDB: 4QI1). Translation/libration/screw (TLS) vibrational motion was applied with TLS groups determined by the TLSMD server (Painter and Merritt, 2006). For details on refinement statistics see Table S2. RMSD values were calculated using PyMOL (PyMOL Molecular Graphics System, version 1.3; Schrödinger), and B factors were extracted from PDB files. All structural figures were prepared using PyMOL. Processing, molecular replacement, and refinement software was from the SBGrid Software Consortium (Morin et al., 2013). ACCESSION NUMBERS The atomic coordinates have been deposited in the PDB, under accession codes PDB: 5ITE (in meso crystal structure of HwBR from OG micelles) and PDB: 5ITC (in meso crystal structure of HwBR from SMA nanodiscs). SUPPLEMENTAL INFORMATION Supplemental Information includes Supplemental Experimental Procedures, three figures, two tables, one note and can be found with this article online at Structure 25, , February 7, 2017

9 AUTHOR CONTRIBUTIONS J.B. and O.P.E. designed the research and wrote the manuscript. J.B. performed the research. J.B. and B.T.E. solved the crystal structures, and B.T.E. commented on the manuscript. ACKNOWLEDGMENTS We are grateful to Heiko Heerklotz (University of Toronto) for access to DLS. We thank Johannes Heidemann and Wei-Lin Ou (University of Toronto) for providing plasmid HwBR-pET28b. We also thank Hazeem Alhalabi and Aidin Balo (both University of Toronto) for assistance with LCP and help with Phenix, respectively. This research used resources of the Advanced Photon Source (APS), a US Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under contract no. DE-AC02-06CH We particularly thank the staff at beamlines 17-ID-B and 23-ID-B/D as well as X-CHIP Technologies Inc. (Toronto). We are also grateful to Onno Looijmans and Niels O. Riekerink (Polyscope) for supplying us with SMA copolymer. For fruitful discussions and carefully reading the manuscript we thank Sebastian Fiedler, Henrike M uller-werkmeister, and Emil F. Pai (all University of Toronto). This work was supported by a Research Fellowship from the German Research Foundation (DFG; to J.B.; BR 5124/1-1) and by the Canada Excellence Research Chairs program (to O.P.E.). O.P.E. holds the Anne and Max Tanenbaum Chair in Neuroscience at the University of Toronto. Received: July 27, 2016 Revised: September 20, 2016 Accepted: December 12, 2016 Published: January 12, 2017 REFERENCES Abla, M., Unger, S., Keller, S., Bonneté, F., Ebel, C., Pucci, B., Breyton, C., and Durand, G. (2015). Micellar and biochemical properties of a propyl-ended fluorinated surfactant designed for membrane-protein study. J. Colloid Interf. Sci. 445, Adams, P.D., Afonine, P.V., Bunkoczi, G., Chen, V.B., Davis, I.W., Echols, N., Head, J.J., Hung, L.W., Kapral, G.J., Grosse-Kunstleve, R.W., et al. (2010). 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