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Membrane proteins play a tremendously important role in cell physiology and serve as a target for an increasing number of drugs. Structural information is key to understanding their function and for developing new strategies for combating disease. However, the complex physical chemistry associated with membrane proteins has made them more difficult to study than their soluble cousins. Electron crystallography has historically been a successful method for solving membrane protein structures and has the advantage of providing the natural environment of a lipid membrane. Specifically, when membrane proteins form two-dimensional arrays within a lipid bilayer, images and diffraction can be recorded by electron microscopy. The corresponding data can be combined to produce a three-dimensional reconstruction which, under favorable conditions, can extend to atomic resolution. Like X-ray crystallography, the quality of the structures are very much dependent on the order and size of the crystals. However, unlike X-ray crystallography, high-throughput methods for screening crystallization trials for electron crystallography are not in general use. In this chapter, we describe two alternative and potentially complementary methods for high-throughput screening of membrane protein crystallization within the lipid bilayer. The first method relies on the conventional use of dialysis for removing detergent and thus reconstituting the bilayer; an array of dialysis wells in the standard 96-well format allows the use of a liquid-handling robot and greatly increases throughput. The second method relies on detergent complexation by cyclodextrin; a specialized pipetting robot has been designed not only to titrate cyclodextrin, but to use light scattering to monitor the reconstitution process. In addition, the use of liquid-handling robots for making negatively stained grids and methods for automatically imaging samples in the electron microscope are described.
Electron microscopy (EM) has made a significant contribution to our understanding of membrane protein structure through the application of electron crystallography (1–3). As with X-ray crystallography, the formation of suitable crystals is the first and often biggest hurdle to overcome (4). Crystallographic methods cannot be applied without crystals and their quality is primarily responsible for the resolution of the final structure. The number of structures solved by X-ray crystallography has experienced exponential growth in the last two decades. Although membrane proteins have lagged behind their soluble counterparts, recent successes show a marked acceleration not only in numbers of structures, but also in their biological impact (5). Much of this success is attributable to automation, which allows X-ray crystallographers to implement high-throughput approaches at various stages of the crystallization pipeline. Specifically, automation is employed for screening genetic constructs for expression, screening of detergents for protein stability and, of course, screening tens of thousands of conditions for producing well ordered crystals (6; 7). With regard to EM, methods for automation are routinely employed for collecting image tilt series for tomographic reconstruction (8–12) and for collecting images to include in single particle reconstructions (13–16). In addition, prototypical robotic systems have been reported for exchanging samples in the electron microscope (17–19). However, relatively little attention has been paid to automating the process of forming two-dimensional (2D) arrays of membrane proteins within the lipid bilayer, so-called 2D crystals. Such regular assemblies are amenable to atomic scale resolution assessment by electron crystallography and they yield the structure of membrane proteins within their native lipid bilayer environment, thus promoting a physiologically relevant conformation. High-throughput automation of 2D crystallization is critical to more wide-spread application of electron crystallography. In this review, we will describe recent developments in high-throughput 2D crystallization employing both detergent dialysis and detergent complexation with cyclodextrin. In addition, we describe facilities necessary for imaging these large-scale 2D crystallization screens in the electron microscope.
As with 3D crystallization of detergent-solubilized protein for X-ray crystallography, formation of 2D arrays within a lipid bilayer is favored by a homogeneous starting solution, in which the protein adopts a single oligomeric state and the mixed micelles of protein, detergent and lipid are monodisperse. Detergents for promoting monodispersity and stability can be screened using an FPLC system equipped with a size exclusion chromatography column. A protein solution that is well behaved will form a single Gaussian-shaped peak during elution from the column. The presence of additional peaks in the void volume, or the presence of multiple or asymmetric peaks in the elution volume indicates that the protein micelles are not monodisperse, either due to the presence of multiple oligomeric states or aggregation. After using chromatography to optimize the biochemical parameters of a preparation, it is also useful to examine the sample by negative stain EM to further verify its monodispersity and stability over time.
Prior to forming 2D arrays, the membrane protein must be integrated into a lipid bilayer. Although, conceptually, a detergent-solubilized protein could be introduced into a preformed lipid bilayer, the more general procedure begins with a fully detergent-solubilized solution of lipid and protein; a variety of methods are then used to reconstitute the lipid bilayer under conditions that favor insertion of the protein and formation of an ordered 2D array. More specifically, this solubilized state is maintained by an excess of detergent, which accommodates both lipid and protein molecules in the micellar state. When the detergent is removed from the solution--e.g., by dialysis, by complexation or by dilution--the lipid is forced to self-associate and form bilayers. The protein has the choice either of inserting into this bilayer, or of aggregating into an insoluble precipitate. Formation of ordered arrays within the bilayer requires a regular network of protein-protein contacts that is generally influenced by the nature of the lipids and by the components of the aqueous phase. Although the definition of suitable conditions is largely empirical, there are several general principles that govern the reconstitution process. (1) Membrane proteins generally do not tolerate long periods at high detergent concentrations and, furthermore, this fully solubilized state has no productive role in the reconstitution process. (2) The transition from the micellar state to the bilayer state is often critical in producing well ordered crystals and this transition should be made relatively slowly. (3) Temperature is often critical, inducing denaturation of detergent-solubilized protein if too high and preventing necessary reorganization of protein within the reconstituted bilayer if too low (i.e., below the liquid to gel phase transition temperature). (4) Although overall protein concentration is less important, the lipid-to-protein ratio is key to crystallization. Although reconstitution can be readily achieved at high lipid-to-protein ratios (and may be optimal for functional characterization), proteins will have a harder time making the necessary network of interactions in the presence of excess lipid. On the other hand, insufficient amounts of lipid will lead to protein aggregation and precipitation.
Prior to screening a membrane protein for crystallization, it is essential to characterize the relative amounts of protein, detergent and lipid in the starting solution. This information allows one to accurately and reproducibly control important parameters such as the lipid-to-protein ratio and the rate of detergent removal. In the following methods, we present two alternative approaches for removing the detergent, namely by dialysis and by complexation with cyclodextrin. Both of these techniques are amenable to parallelization, though some special facilities are required. Specifically, a 96-well dialysis chamber has been devised, which is compatible with the use of a liquid-handling robot or multi-channel pipettor. For cyclodextrins, a pipetting robot has been designed and built to add nanoliter quantities of cyclodextrin stock solutions systematically to commercially available 96-well microtiter plates. This so-called 2DX robot is also equipped with a level sensor, to allow compensation for evaporation, a shaker to ensure mixing, a temperature controller, and a light scattering detector to follow the reconstitution process. All of these parameters are displayed in real time by the control software. Alternate approaches for crystallization are dilution of the detergent below its cmc (23) or addition of polystyrene beads (BioBeads) to adsorb the detergent (24). Although the former is suitable for parallelization, it’s disadvantage is the inevitable dilution of the protein, which becomes impractical with long-chain (low cmc) detergents. Whereas good results have been obtained with BioBeads, this method is not amenable to automation, given the difficulty in handling the beads.
In this section, we present methods for determining protein concentration using SDS-PAGE, detergent concentration using contact angle measurements, and lipid concentration using TLC. Although there are easier, faster ways to determine protein concentration (i.e. absorbance at 280 nm, Lowry assay, Bradford Assay), these are often inaccurate and are sometimes influenced by the detergent present in the solution. We therefore recommend SDS-PAGE to compare the staining of the purified protein relative to known amounts of BSA. This method removes the variability due to detergent, requires only minimal amounts of sample. Furthermore, the Coomassie-stained protein band can be excised from the gel and sent to a protein sequencing facility to confirm identity. It should be noted that the Coomassie stain used for SDS-PAGE does not stain all proteins equally, so the comparison of staining intensity relative to BSA cannot be assumed to represent an exact measure of concentration. Moreover, SDS-PAGE is time consuming and may not be the best choice for unstable proteins. In these cases it may be necessary to use one of the aforementioned spectroscopic techniques, perhaps after calibrating them relative to SDS-PAGE. The NanoDrop 2000c spectrophotometer is an excellent alternative given its extremely small sample size.
Purified membrane proteins may have a poorly defined detergent concentration due to the use of a concentrator, which often retains detergent and increases its concentration in an unpredictable way. Although a concentrator with a molecular weight cutoff of 50 kDa will remove short- and medium-chain detergents (e.g., OG and DM) together with the filtrate, long-chain detergents are generally retained with the protein with molecular weight cutoffs up to 100 kDa, which may be larger than the expected micelle size. In this case, detergent concentration can be determined from the shape of droplets. This shape is governed by surface tension. Detergent molecules that line the air/water interface decrease the surface tension and cause the drop to spread. The contact angle between the drop and the supporting surface decreases monotonically with increasing detergent concentration up to the cmc. This behavior is characteristic for all detergents and needs to be calibrated (20). Above the cmc, the concentration of free detergent molecules remains unchanged, so there are no further changes in drop shape. Thus, solutions with higher detergent concentrations, such as those used to purify proteins, must be diluted to bring the detergent below the cmc. This may cause the protein to precipitate, but this precipitate can be removed by centrifugation.
It has frequently been observed that lipids co-purify with membrane proteins. Also, addition of exogenous lipid is a common way to optimize membrane protein stability during purification. TLC is a well-established method both to quantify the total amount of lipid and to identify different lipid species. Specifically, if the appropriate standard lipids are run together with the protein sample, it is possible to fully characterize the lipid composition in the solution.
The dialysis and cyclodextrin methods represent two complimentary approaches to crystallization. On the one hand, the ability to dictate the rate of cyclodextrin addition can be used to precisely control the rate of detergent removal and consequent reconstitution. It is reasonable to assume that the transition between the micellar phase and the bilayer phase is critical to the crystallization process, and manipulation of cyclodextrin addition offers the opportunity to explore this transition. On the other hand, dialysis has a longer track record in reconstitution and membrane protein crystallization. Although the level of control is somewhat limited, the resulting solutions do not have high levels of cyclodextrin present, simplifying the preparation of samples for electron microscopy. With either approach, differences in the physical chemistry of detergent removal can be expected to produce different results and it is impossible to know a priori which will yield the best crystals for a given protein target. In the spirit assessing the affect of all possible parameters, it may be useful to attempt both methods of crystallization. This is analogous to empirical testing of different 3D crystallization methods for X-ray crystallography (e.g., hanging drop, sitting drop, batch, lipidic cubic phase) in order to obtain the best possible result.
A combination of these two methods may also be a productive avenue to explore. In particular, the addition of cyclodextrin to the dialysis buffer has the potential to accelerate and to provide finer control over dialysis rates. Dialysis rates are fundamentally limited by the relatively low concentration of monomeric detergent molecules, which are the only species that can equilibrate across the dialysis membrane. Detergent micelles typically contain ~100 molecules and are therefore too large to move across this membrane. The limitation is particularly acute for long-chain detergents, which have a very low cmc. However, both cyclodextrin and the cyclodextrin-detergent complexes are able to migrate across the dialysis membrane, thus effectively increasing the pool of monomeric detergent molecules that can be equilibrated and thus removed by dialysis. Preliminary analysis has shown cyclodextrin to accelerate the removal of DDM greatly, achieving complete detergent removal in only 4 days, compared with the typical 2 weeks in the absence of cyclodextrin. The overall cyclodextrin concentration remains relatively low throughout the process and can be completely eliminated by simply omitting cyclodextrin in the last change of dialysis buffer. Because this buffer is frequently changed throughout the procedure, one can maintain a certain control over the rate of detergent removal, for example, pausing when the protein solution reaches the transition from micellar to bilayer phases. Also, a bolus of cyclodextran can be added to the starting protein solution to establish a starting condition that is relatively close to the transition, thus minimizing the unproductive period that a protein spends in the micellar state. This may be particularly important for unstable membrane proteins.
The authors would like to acknowledge a number of individuals who have developed the methodologies discussed in this chapter. In particular, Martin Vink, Changki Kim and Minghui Hu, Thomas Kaufmann, Ioan Iacovache, Nicolas Coudray Hervé Rémigy and many others were responsible for designing and implementing the devices and protocols described in this chapter. The authors are indebted to them for their efforts. The authors belong to the Transcontinental EM Initiative for Membrane Protein Structure, which is a center for membrane protein structure determination funded by the NIH Protein Structure Initiative under grant U54GM094598. More information about this center can be found at http://temimps.nysbc.org. Additional research support was provided by NIH grants R01GM081817 (D.S. and I.U.-B.), R01GM095747 (D.S. and I.U.-B.), R01GM079233 (T.G.), by the NSF CAREER award MCB-0546087 (I.U.-B.) and by the American Diabetes Association Career Development award 1-09-CD-05 (T.G.). T.G. is a Howard Hughes Medical Institute Early Career Scientist.
1 Ideally, protein is used immediately after purification without freezing, storage or concentration. By freezing or otherwise storing the protein, the possibility of denaturation or aggregation increases. Concentrators typically increase the detergent concentration in an unpredictable way, making it difficult to precisely define the starting point for crystallization.
2 The cmc is defined as the concentration at which micelles start to form. Below this concentration, all detergent molecules are monomeric in solution. Above this concentration, there is a mixture of monomeric detergent molecules and micelles, which are in dynamic equilibrium. Above the cmc, the concentration of monomers remains constant at the cmc. The cmc is characteristic for each detergent, depending on its chemical composition, and is affected by physical/chemical parameters such as the presence of solutes and temperature.
3 This selection of lipids has hydrocarbon chain lengths of 14 (myristol), 16 (palmitoyl) and 18 (oleoyl). The latter has one unsaturated bond, whereas the former are both fully saturated. The head groups range from the zwitterionic phosphatidylcholine and phostatidyl ethanolamine, to the negatively charged phosphatidylglycerol and phosphatidic acid. Although this is a good starting set of lipids, in many cases it is also worth using other membrane components, such as cardiolipin, cholesterol and spingolipids, or using lipid extracts from the relevant organism or tissue, such as liver, heart, brain E. coli, or yeast.
4 Parafilm is the standard substrate reported by Kaufmann et al. (20) that generally works well. Teflon tape (e.g., for pipe fittings) is an alternate choice of substrate and may be suitable for short-chain detergents. Parafilm has the advantage of maximizing changes in the contact angle as a function of concentration. Teflon is more hydrophobic and thus limits the extreme flatness of drops produced by short-chain detergents (e.g. OG) as their concentration approaches the cmc. With some software, it is difficult to determine the contact angle for very flat drops and measurements may be therefore be more reproducible using Teflon tape.
5 Although contact angle is the conventional and most physically rigorous measure of surface tension, the ratio of drop width and height also produces reliable results, at least for small drops (e.g., 20 μl where the shape is well approximated by an ellipse). The programs Xdroptrace and DropBox both fit the drop shape with an ellipse and calculate either the width/height ratio (based on the major and minor axes) or the contact angle (based on the intersection with the planar substrate). The measurement of contact angle is very sensitive to the precise position of the substrate and the axial ratio may therefore be more robust in practice. Both Xdroptrace can be downloaded from http://temimps.nysbc.org.
6 If the concentration of lipid in the protein solution is low, then the lipids may be readily extracted from the solution with chloroform. After separating the organic phase from the aqueous phase, the chloroform can be evaporated and the resulting lipid film resolubilized in a smaller amount of chloroform prior to running the TLC. Extraction into an organic phase is also a method for separating the lipid from the detergent, which will be abundant and perhaps interfere with the signal from the lipid. Although nonionic detergents partition between the aqueous and organic phases, multiple washings of the organic phase with water will greatly reduce the amount of detergent present; lipid has a negligible solubility in water and will remain almost completely in the organic phase.
7 It is judicious to include a small amount of butylated hydroxytoluene (BHT) as a scavenger of free radicals and therefore as a mechanism to minimize lipid oxidation. Prepare a stock solution of 20% BHT in ethanol and add a sufficient amount to obtain 0.2% BHT in the final aqueous stock solution of lipid.
8 It is possible to bring lipid into the aqueous phase in the absence of detergent, but the lipid will form very heterogeneous multivesicular structures that may prove difficult to solubilize at a later stage. A more homogeneous, pure lipid solution can be obtained by sonicating the solution using a probe sonicator (e.g. Branson Sonifier S-250 fitted with a microtip). The effectiveness at producing unilamellar vesicles depends on the nature of the lipid; lipids with a net charge more readily form unilamellar structures, whereas neutral lipids are more difficult to disperse. Sonication puts a significant amount of energy into the solution and can cause chemical changes (oxidation, cleavage of headgroups, creation of lyso-lipids); the solution should therefore be kept cold (on ice and in a cold room) and sonication periods should be minimal (1–3 minutes with 50% duty cycle and with breaks to allow the solution to remain cold). Since a detergent-solubilized, homogeneous solution is required to start crystallization, it therefore makes sense to directly solubilize the lipid film with detergent and avoid these potential problems, but the ability of a given detergent to solubilize the lipid also depends on the physical state of the lipid.
9 To remove the EM grids from the water surface, layer the newspaper on top of the EM grids, allow the paper to absorb a bit of water and then peel the paper off of the surface, thus removing the EM grids as a sandwich between the newspaper and the plastic film. Place the sandwich into a petri dish with the plastic facing upwards and allow to air dry (30).
10Depending on the apparatus, either a carbon rod (graphite or amorphous carbon) or a carbon thread can be used for evaporation. For high resolution work, the flatness and conductivity of the carbon support is critical (31). For evaluating negatively stained samples, however, one simply requires a uniform carbon film that provides a reasonable proportion of unbroken grid squares (the broken squares can be identified by automated imaging programs and easily avoided). The underlying plastic film is helpful in this regard. It is also important to control the current during carbon evaporation in order to avoid sparking, which produces an inhomogeneities in the resulting film. Also, many investigators evaporate several thin layers of carbon in order to improve strength.
11 An optional step in EM grid preparation is to remove the plastic film prior to glow discharge. This improves the clarity of high magnification images and also reduces residual stickiness that results from the ragged edges of the plastic film at the periphery of the grid. However, this step is not easily automated and represents an interruption in the workflow. To remove the plastic, place several pieces of filter paper into a glass petri dish. Saturate the filter paper with amyl amine. Place EM grids onto the filter paper with carbon side facing upwards. Cover the petri dish and allow grids to incubate for 15–30 min in a chemical fume hood). Remove the grids and place on a fresh piece of filter paper for glow discharge. An alternative method for producing carbon film to is evaporate carbon onto a freshly cleaved mica surface and then to float this carbon onto a water surface. This method, also less amenable to high throughput methods due to fragility in the resulting film, is described by Stokes and Ubarretxena (30).
12 The choice between blotting and aspiration will depend on the quality of the samples as assessed by electron microscopy. The goal is to obtain an even, thin layer of solution across the entire EM grid, which after drying encases all of the samples in a even layer of negative stain.