Crystallizing Membrane Proteins Using Lipidic Mesophases

Keywords: thin layer chromatography, polarized light microscopy, small-angle X-ray scattering, fluorescence, ligand binding, monoolein, BtuB, OpcA, LH2, GPCR

An overview of protein crystallization methods

Membrane proteins can be crystallized for use in macromolecular crystallography (MX) by a variety of methodologies. These have been divided into two major groupings, the so-called in surfo and the bilayer methods³.

In meso crystallogenesis

In meso crystallogenesis takes place in and from a bicontinuous lipidic mesophase. It is important to appreciate that such mesophases are dynamic structures that respond to both compositional and environmental conditions in a sample history-dependent way. In the simplest case, composition refers to the relative amount of lipid and water in the system. However, under conditions of crystallogenesis, the system actually consists of an array of different components that can include protein, detergent, buffer, salt, hydrogen ions, polymers and small molecule additives. Any and all of these can influence lipid phase behavior and it is likely that the manner in which they do so influences the outcome of a particular crystallogenesis trial. It is important therefore to be familiar with the basic phase behavior of the hosting lipid with a view to understanding what is happening at a molecular, structural and a mesophase level at the various stages in crystallogenesis. For this reason, a very brief introduction will be given here to the temperature-composition phase diagram of monoolein, the most successful lipid used to date for in meso crystallogenesis¹³ (Figure 1). At its simplest, the phase diagram is a concise summary of phase propensity as water content and temperature are changed (see refs. ¹⁴ and ¹⁵, for useful background) The focus here is on the 20 °C isotherm, which is where the bulk of in meso crystallization has been performed (Figure 1).

As noted, crystallogenesis begins with the cubic mesophase. The latter is accessed in the monoolein/water system at 20 °C at an overall composition of about 40 % (wt/wt) water (A in Figure 1). This corresponds to the cubic phase of space group type Pn3m (cartoon representation of the phase is shown in Figure 1). When additional water is added beyond the hydration limit of ~40 % (wt/wt) water, it appears as a second phase and a two-phase system forms (B in Figure 1). With reference to the crystallization trial, this corresponds to the cubic phase bolus sitting in an excess of precipitant solution. Moving along the 20 °C isotherm in the direction of reduced hydration, the next phase encountered after the cubic-Pn3m phase is a second cubic phase, this time of space group type Ia3d (C in Figure 1). A continued reduction in water content triggers formation of the lamellar liquid crystalline or L_α phase (D in Figure 1). The latter consists of planar lipid bilayers stacked atop one another and separated from each other by a layer of water. A further reduction in water gives rise to the lamellar crystalline or Lc phase which is not liquid crystalline but a true solid (E in Figure 1). Depending on how the lipid was treated thermally prior to manipulation, the system can at this point access what is referred to as the fluid isotropic phase which, like the Lc phase, is not a mesophase but a true liquid. One final point of note regarding the phase diagram is that the excess hydration boundary shifts from about 50 % (wt/wt) water at 0 °C to 25 % (wt/wt) water at 85 °C. This temperature-induced shedding of water by the fully hydrated cubic mesophase is a useful diagnostic referred to later in this Protocol.

That the method works and that it is a generally applicable method is no longer in doubt as evidenced by the statistics to be presented below. Less certain however is the mechanism by which it occurs. A hypothesis has been advanced for how the method works³ and convincing experimental evidence is available in support of the model (Figure 2). The proposed mechanism begins with the target protein being solubilized in detergent from the parent biomembrane or from inclusion bodies. It is then purified using standard methodologies that include gradient centrifugation¹⁶ and affinity¹⁷, ion exchange¹⁸ and size-exclusion chromatographies¹⁹. The purified, concentrated protein in detergent solution is combined with lipid that, when fully hydrated, forms a viscous and sticky cubic mesophase. At this stage, the protein is considered to have been reconstituted into the bilayer of the cubic phase in a native and active conformation. The system is now destabilized by overlaying the mesophase with an excess of precipitant solution. The latter does not normally dissolve the cubic phase but exists as a separate liquid from which components diffuse into the mesophase bolus. The bolus can be considered a molecular sponge with the lipid bilayer corresponding to the fabric of the sponge and the aqueous channels that permeate the mesophase corresponding to pores in the sponge. Components in the precipitant solution diffuse into the mesophase and trigger a series of phase transitions and separations, one of which leads to nucleation and growth of a protein crystal.

Limitations of the in meso method

The in meso method was introduced over a decade ago²⁰. Thus far, it is credited with 54 entries in the Protein Data Bank (see ref. 21; www.mpdb.ul.ie), which corresponds to about 10 % of the integral membrane protein structures in the public domain. Our sense is that the yield of structures is not as high as it might be because the method has one major challenge; the cubic phase is extremely viscous (akin to a thick toothpaste) and sticky. As a result it is not easy to handle with the tools commonly found in a biochemistry or a structural biology laboratory. Many have tried to use the method but have abandoned it in frustration because the material at the heart of the method, the cubic phase, was found to be difficult to prepare and to dispense. The purpose of this Protocol is to explain how to work with such mesophases and to provide simple and clear, step-by-step instructions for performing in meso crystallogenesis and related studies. The Protocol is based on methods in routine use in the authors’ laboratories and that have been employed in characterizing and determining the high-resolution structures of the light-harvesting complex 2 from R. acidophila²², the outer membrane Vitamin B12 transporter BtuB from E. coli²³, the outer membrane adhesin/invasin OpcA from N. meningitidis²⁴ and the human β₂-adrenergic²⁵ and adenosine A_2A²⁶ G protein-coupled receptors (GPCR). It is recommended that initial trials (Steps 1–35) be performed with lipid and buffer in place of the target protein solution to gain familiarity with the method and with the unusually viscous and sticky nature of the mesophase.

Automation of in meso crystallogenesis

In meso crystallogenesis can be done manually²⁷, or robotically in high-throughput mode²⁸. The former is emphasized here because it can be done inexpensively and with materials and supplies that are readily available. We use the manual approach for optimization trials that involve small numbers of samples. For high-throughput applications we have built an in meso crystallization robot (Figure 3) and a brief description of its use is included in Box 1. The robot offers distinct advantages such as throughput, uniformity, and the ability to perform the set up under what might be considered hostile conditions such as in darkness and at low temperatures. It can also operate with extremely small volumes of protein (in the picoliter range) responding to the need to miniaturize to conserve valuable protein (and lipid). The steps that must be taken for such low volume setup are described in ref. 29.

Overview of the Procedure

The Protocol is arranged by section using the procedural sequence followed in the laboratory for a typical crystallogenesis trial (Figure 4). Crystallization plates are first prepared, and then the lipid-protein mixing device is loaded and used to homogenize components. This enables the protein to reconstitute into the bilayer of the spontaneously formed cubic mesophase. Wells in crystallization plates are loaded with the protein-containing mesophase, which is then bathed in precipitant solution. The plates are sealed creating a glass sandwich of the mesophase which offers unparalleled optical quality for subsequent imaging. Incubation and interrogation by normal and polarized light microscopy follow. Hits, when identified, are followed up with rounds of optimization that involve an exploration of incubation temperature, and precipitant and mesophase composition. The latter refers to lipids that create the hosting mesophase and to lipid additives. The next stage in the process involves harvesting crystals and is followed by diffraction measurements. Because in meso-grown crystal can be small and in an opaque medium upon cryo-cooling use is typically made of an X-ray synchrotron minibeam to locate and to orient the crystal ahead of data collection. The latter is not covered in this Protocol. In addition to laying out the procedures for performing in meso crystallogenesis and for crystal harvesting, the Protocol includes methods that form part of the pre-screening exercise, which is performed to minimize wasted time and effort with systems defective to begin with and that will not yield diffraction quality crystals. These include: methods used to characterize lipid purity and phase properties, including thin layer chromatography, small-angle X-ray scattering and polarized light microscopy (Box 2) and; a description of how the conformation and activity of the protein reconstituted in the cubic phase is evaluated by a variety of spectroscopies including UV-visible, fluorescence, and circular dichroism (Box 3). Pre-screening and optimization should be performed by carrying out steps 1–13 of the main Procedure, followed by Boxes 2 and 3 taking care to set up the appropriate controls at relevant steps. When happy with the results of the analyses, carry out the crystallization experiment by repeating the Procedure from step 1 using the optimized conditions, continuing directly with Step 15.

REAGENTS

Monoolein (1-oleoyl-rac-glycerol) (Sigma, or Nu Chek) (Refer to Box 2 for methods to characterize lipid purity and phase properties)

CRITICAL. The monoacylglycerols used are unsaturated and can undergo oxidation. It is advised to always store stock lipid in the dark at as low a temperature as possible (−20 or −80 °C) in an inert atmosphere of nitrogen or preferably argon gas.

Phospholipids, cholesterol (Avanti Polar Lipids) (Refer to Box 2 for methods to characterize lipid purity and phase properties)

Crystallization screens (Hampton Research, Molecular Dimensions, Jena Biosciences, Qiagen, Emerald Biosystems)

Chloroform. CAUTION toxic/irritant. Wear protective gloves and perform all manipulations under a fume hood.

Methanol. CAUTION flammable/toxic. Wear protective gloves and perform all manipulations under a fume hood.

Silanizing agent (for example, AquaSil from the Pierce Division of Thermo Fisher Scientific, Cat. No. 42799). CAUTION flammable/irritant. Wear protective gloves and perform all manipulations under a fume hood.

EQUIPMENT

Access to a microwave oven, hot running water or a small heating block that is stable at any where from 40 to 60 °C is needed to melt the lipid.

Analytical balance with sub-milligram sensitivity.

Assortment of Eppendorf tubes and rack.

Eppendorf microcentrifuge capable of generating ~14,000 g.

Ultrasonic humidifier (optional).

One 20 – 100 μL, one 1 – 10 μL, and one 0.2 – 2 μL adjustable pipette with disposable tips. We typically use Pipetman pipets.

10 μL Hamilton gas-tight syringe with a removable needle (Hamilton, Cat. No. 80065).

Two 100 μL Hamilton gas-tight syringes with a removable needle (Hamilton, Cat. No. 81065).

25 μL Hamilton gas-tight syringe with a removable needle (Hamilton, Cat. No. 80265).

Repeating dispenser (Hamilton, Cat. No. 83700). Optionally, the dispenser can be modified, as described in ref. 30 to reduce the volume of cubic phase dispensed with a 10 μL syringe from 200 to ~70 nL.

Two removable needles (gauge 22, Hamilton, Cat. No. 7770-020) and two RN nuts (Hamilton, Cat. No. 30902) to make a syringe coupler (see Equipment Setup section). Alternatively, the syringe coupler can be purchased commercially as part of the Cubic LCP kit (Emerald Biosystems, Cat. No. EBS-LCP-2). However, the critical threaded parts of this commercial coupler are made of plastic, whereas the original is made of steel and is extremely durable. We have not tested the commercial coupler and are not in a position to comment on its performance and/or suitability.

Short (0.375 inch), flat-tipped needle (point style 3, gauge 26, Hamilton, Cat. No. 7804-03)

Optional. A desk lamp and a magnifying lens, to aid in viewing syringes, coupler and plates during loading and dispensing, are useful but not essential. A combined lamp/magnifying lens unit can also be used.

Microscope slides, 1 inch × 3 inch (GoldSeal, Cat. No. 3010)

Microscope cover slips, 1 inch × 1 inch (Fisher Scientific, Cat. No. 12-548C)

Perforated double stick spacer (3M 9500PC double stick tape, 0.140 mm thick, 5 mm diameter holes, 27 holes total, 7 mm spacing between holes, overall size 1 inch × 3 inch. The arrangement of holes in the 27-well plate is shown in Figure 5 (see also ref. 27). Spacers with punched holes can be obtained from a die-cutting company such as Saunders Corporation (http://saunderscorp.com)).

Two wash bottles with fine tips. One filled with milli-Q or distilled water, the other with methanol. These are used to clean the syringes, needles and coupler. Methanol is used first, then water.

Gloves to protect hands whilst working with organic solvents.

Pressure-regulated air or nitrogen gas with a hose and a fine tip for drying syringes, needles, ferrules, and coupler. A can of pressurized air works well.

Parafilm, 1 cm × 1 cm and 1 inch × 1 inch squares.

Hand held roller or brayer (Fisher Scientific, Cat. No. 50820937)

Tools for Harvesting Crystals from Glass In Meso Crystallization Plates:

• ○ Hampton glass cutting stone, broken to present a fresh, sharp edge (Hampton Research, Cat. No. HR4-334).

• ○ Fine point tweezers (Ted Pella, Cat. No. 510).

• ○ Probing tool with a sharp point angled at 45 degrees (Ted Pella, Cat. No. 13650).

• ○ In meso crystallization plate with protein crystals.

• ○ Precipitant solution for the condition that supports crystal growth.

• ○ Stereo microscope with crossed polarizer capability, magnification range 20 – 100x and with a large operating distance (>60 mm) for easy access to plate and for ease in harvesting crystals

• ○ Pipet with tip set to 5 μL for transferring the precipitant solution.

• ○ Magnetic wand (Hampton Research, Cat. No. HR4-729).

• ○ Covered dewar (Hampton Research, Cat. No. HR4-673) filled with liquid nitrogen.

• ○ Storage or shipping dewar (Taylor-Wharton).

• ○ Magnetic caps. Standard and ALS-style caps are available from Hampton Research; SPINE-style caps are available from Molecular Dimensions and Hampton Research. Check with your beamline host for acceptable cap styles.

• ○ Cryoloops. For harvesting small crystals from lipidic cubic phase we recommend MicroMounts with 20 to 50 μm aperture, available from MiTeGen (Cat. No. M1-Lxx-20, M1-Lxx-30, M1-Lxx-50) (https://mitegen.com/mic_catalog.php?c=MicroMounts). Alternatively, nylon cryoloops can be obtained from Hampton Research or LithoLoops can be purchased from Molecular Dimensions. Check with your beamline host for acceptable pin lengths.

• ○ Super Glue (Hampton Research cat. No. HR4-316) or an Epoxy (Hampton Research cat. No. HR4-318)

• ○ OPTIONAL. Puck to house multiple mounted crystals. These are typically used with automounters. However, some pucks, such as ALS-style pucks, are also convenient for manual mounting (ALS-style pucks and accessories can be purchased from Boyd Technologies, www.boydtechnologies.com; SSRL cassette and Unipucks can be purchased from Crystal Positioning System, www.crystalpositioningsystems.com; Rigaku-style pucks can be purchased from Rigaku; EMBL/ESRF pucks and accessories are available from Molecular Dimensions)). Check with your beamline host for puck styles that are compatible with their automounter.

• ○ OPTIONAL. Cryovials. For manual mounting (Hampton Research cat# HR4-731). Some automounters, such as Rigaku and EMBL/ESRF, require cryovials (CrystalCap HT from Hampton Research; SPINE style magnetic cryovials from Molecular Dimensions).

• ○ OPTIONAL. CrystalCap holder (Hampton Research cat# HR4-707) is used to hold cryovials in the dewar during crystal harvesting.

• ○ OPTIONAL. CryoCane (Hampton Research cat# HR4-709 or HR4-711) and CryoSleeve (Hampton Research cat# HR4-708) are used for storing and transporting mounted crystals in Cryovials.

REAGENT SETUP

CRITICAL: Preformulated buffers, precipitants (salts, polymers, volatile and nonvolatile organic solvents), detergents, surfactants, additives, reducing agents and ligands required for the solutions below are available commercially (see web sites for Hampton Research, Molecular Dimensions, Jena Biosciences, Qiagen, Emerald Biosystems, Sigma-Aldrich, etc.). These reagents are used to prepare various buffers and precipitant cocktails. The different sources vary in the range of ingredients, concentrations and volumes provided. Refer to manufacturer instructions for maximum storage time and preferred storage conditions of light and temperature.

CRITICAL Reproducible crystallogenesis benefits from the use of materials of the highest and most uniform quality and purity.

CAUTION Many of the reagents used in crystallogenesis are toxic and instructions for handling, as described on the relevant Materials Safety and Data Sheets, should be followed.

EQUIPMENT SETUP

Loading the Lipid Mixer

TIMING 10 min.

1) Remove the lipid, monoolein, from the freezer and allow it to adjust to room temperature (~20 °C). CRITICAL STEP. Monoolein, as well as other monoacylglycerol lipids, are hygroscopic. Therefore, allow for full equilibration to room temperature before opening the container.

2) Disconnect the needle and remove the Teflon ferrule from two 100 μL Hamilton gas-tight syringes.

3) Weigh both syringes with plungers in place.

4) Remove the plunger from one of the syringes.

5) With a fine tipped spatula, work about 10–30 mg (approximately 10–30 μL, density of monoolein at 20 °C is 0.942 g/cm³, CRC Handbook of Chemistry and Physics) of the waxy lipid (or lipid mixture, see Box 4 and Box 5) into the open end of the syringe. Return the plunger to the barrel and place the syringe in an oven at about 37 °C to melt the lipid. Alternatively, the lipid can be melted before loading into the syringe. In this case, a 20 or 200 μL pipet with a standard disposable tip is used to remove 10–30 μL of the molten lipid and to place it in the barrel of the syringe (plunger removed) from the plunger end. Return the plunger to the barrel, at which point its Teflon tip will make contact with the molten lipid. With the syringe held vertically, barrel termination end up, move the plunger slowly up the barrel along with the molten lipid; this will usually lead to air locks and bubbles being removed from the lipid. Ideally, one ends up at this stage with a continuous volume of molten lipid in direct contact with the Teflon tip of the plunger. In this case, the volume of lipid in the syringe can be read directly from the μL markings on the barrel. Given that monoolein has a density of 0.942 g/mL (CRC Handbook of Chemistry and Physics), the weight of lipid in the syringe can be calculated from the recorded volume. Alternatively, the loaded syringe can be reweighed and the lipid mass obtained by taring.

CRITICAL STEP. Incubate the lipid for just long enough to melt the lipid for easy handling. Extensive heating can lead to lipid degradation.

6) Attach the coupler, with both ferrules in place, to the open end of the lipid-loaded syringe. Carefully screw the coupler into position. CRITICAL STEP. The coupler must be tightened to a degree that the syringe contents do not leak under the pressure of mechanical mixing but not over tightened which can deform the ferrule and/or break the glued seal between the glass barrel and the steel barrel termination resulting in leakage during mixing.

7) Use the plunger to force the molten lipid up the barrel and slowly and gently into the narrow bore needle at the core of the coupler. If the needle is slightly overfilled, the lipid will be seen beading out at its open end. By backing up the plunger slightly the excess lipid can be withdrawn into the coupler until it is just flush with the tip of the coupler needle. In this case, the syringe/coupler unit is fully loaded and ready to be combined with the protein solution-loaded syringe at step 12.

8) Calculate the volume of protein solution that can/should be used to form the cubic phase at or close to full hydration. For monoolein at 20 °C, full hydration with water occurs at close to ~40 % (wt/wt) water¹³ (Figure 1). Thus, for example, if the weight of lipid in the lipid-loaded syringe is 21 mg then [(21/3) × 2 = ] 14 mg, corresponding to ~14 μL, of protein solution will be required.

9) Centrifuge the protein solution in a 0.5 mL Eppendorf tube at 14,000 g for 5–10 min at 4 °C to remove large aggregates before setting up crystallization trials. CRITICAL STEP. The amount of detergent in the protein solution used for in meso crystallization trials should be minimized to avoid destabilizing the bicontinuous mesophase (see also discussion in the Troubleshooting Section under Step13, Problem: Cloudy mesophase, Possible reason 4: High concentration of detergent in the protein solution).

10) The second 100 μL Hamilton syringe is used to house the protein solution (or control solution required for pre-screening analyses, see Box 3). Withdraw the plunger in the syringe to the 20 – 30 μL mark and slowly inject 14 μL of protein solution onto the Teflon tip of the plunger using a 25 μL Hamilton gas-tight syringe fitted with a 26s gauge, flat tipped (point style 3) needle. CRITICAL STEP. Every effort should be made to avoid trapping air bubbles in the delivered protein solution because removing them after loading can be difficult given the tendency of such detergent-containing solutions to foam. If moving the plunger up and down in the barrel whilst holding the syringe in a vertical position does not eliminate trapped bubbles, it is best to start again. To this end, aspirate the protein solution from the syringe, transfer it back to an Eppendorf tube and centrifuge as in Step 9.

11) At this stage, the protein solution should exist as a homogenous, clear solution in direct contact with the Teflon tip of the plunger. Determine the volume of the protein solution by reading the μL markings on the syringe barrel which should match the volume loaded initially. This is usually the case. Alternatively, the loaded syringe can be reweighed and the weight of the protein solution calculated by taring. In preparation for combining with the lipid-loaded syringe, carefully inch the protein solution up the barrel to end up flush with its open end. This can be observed by looking down through the barrel termination end. A table lamp and magnifying lens are a great help at this step.

12) The lipid- and protein solution-loaded syringes are now ready to be combined in preparation for mixing. To do this, screw the protein syringe into the open end of the coupler attached to the lipid syringe. Having combined the two syringes by way of the coupler a continuous volume of liquid should exist from lipid in one syringe through the coupler to protein solution in the other syringe. In preparation for mixing, the assembled unit is held with one syringe in the right hand and the other in the left. Torque is applied to both barrels through to the coupler using the fingers and thumb of both hands to ensure that the seal against the ferrules in the coupler remains intact. CRITICAL STEP. Over tightening the assembled mixing device at this stage will damage the ferrules and cause leaking. Under tightening will lead to leaking also. It is a matter of experience, with the inevitable trial and error and occasional loss of sample. It is worthwhile therefore practicing with lipid and water or buffer solution to get a feel for the system before launching into using valuable protein solution.

Mixing Lipid and Protein Solution. Making the Mesophase

TIMING 5 min.

13) To effect mixing, advance the plunger on the protein side of the assembled mixing unit to its limit, with the thumb or index finger driving the protein solution out of the protein syringe, through the coupler and into the lipid syringe. If the unit has been sealed effectively this action will cause an equal and opposite movement of the plunger on the lipid side. The plunger on the lipid side is now used to drive the contents of the lipid syringe back through the coupler and into the protein syringe. The process is repeated many times; occasionally a hundred passages or more are required, to produce an homogenous mesophase.

CRITICAL STEP: At the start of mixing, movement of material back and forth through the coupler can be uneven and, at times, extra force is needed to effect mixing. This is to be expected as different mesophases come and go in the course of the homogenization and hydration process. Refer to the 20 °C isotherm in Figure 1 for the phases that are likely to form in the process. Initial mixing is usually accompanied by the development of a non-uniform cloudiness in the sample and is expected. As homogenization progresses the texture, as sensed by the force needed to move the plungers back and forth, becomes more uniform and characteristic of the viscous cubic phase, as does the visual appearance of the emerging cubic mesophase (which is optically clear).

CRITICAL STEP: If the concentration of the protein solution is low (e.g. less than 5 mg/mL) then using an excess of it at this stage can be useful because the protein will preferentially partition into the lipidic mesophase. In this way, the protein concentration in the actual cubic phase can be increased. In practice, a 2- to 5-fold increase in concentration is achievable depending on the detergent concentration in the initial protein sample. The bulk of the excess (protein-free) buffer that spontaneously separates from the fully hydrated mesophase can be easily removed by uncoupling the mixing and retrieving excess liquid using a separate Hamilton syringe. A small amount of the excess buffer, however, will inevitably become trapped in the mesophase and this will contribute to diluting the precipitant solution added later in the process (Step 32). Therefore, it is advisable to add few microliters of lipid at this stage and to mix to homogeneity to absorb the excess buffer.

CRITICAL STEP If conditions are appropriate and the cubic phase forms, the dispersion should appear optically transparent in the syringe barrel. Thus, the markings on the syringe barrel should be clearly legible through the barrel. In the case of colored proteins, the mesophase will have the color of the protein but will be optically transparent. Very slight cooling of the sample during mixing by placing the syringe mixer for a short time on ice, in a refrigerator, or in the air stream of an air conditioning unit can accelerate homogenization and the achievement of transparency. However, it is important not to over-cool the sample, which can destabilize the cubic mesophase (Figure 1) leading to protein damage.

CRITICAL STEP: Care should be taken to avoid extremely vigorous mixing as this can cause sample temperature to rise due to frictional heating³³. Whilst membrane proteins are usually more stable when reconstituted into a lipid bilayer, as is proposed to happen in the mixing process, high temperatures are not likely to be good for the protein. Mixing comes about as a result of shear that develops as the material moves back and forth through the coupler. It is not known how shear will affect the protein but it is likely that the gentler the mixing and the less shear imposed the better. Accordingly, we recommend that the rate of mixing should not exceed ~1 stroke/s.

<TROUBLESHOOTING>

14) To continue with the crystallization process, proceed directly to step 15. Note that for the initial pre-screening stage, the cubic phase and the conformation and activity of the protein reconstituted in the cubic phase should be evaluated as described in Box 2 and Box 3; when happy with the results of the analyses, fresh cubic phase should be prepared (steps 1–13) before proceeding with step 15.

Setting Up Crystallization Plates

TIMING approximately 10 min per 27-well plate.

15) At this juncture the cubic mesophase has been formed and the protein is reconstituted into the lipid bilayer. The protein-loaded mesophase is now ready for dispensing into individual wells of the crystallization plates (steps 29–33). First however the homogenous mesophase must be transferred to a 10 μL Hamilton syringe (RN type) mounted in a repeating dispenser (Figure 8). Begin this process by coupling the syringe to the repeating dispenser, as described in steps 16–21 below. Then load the syringe, as described in steps 22–28, with the cubic mesophase sample (prepared in steps 1–13).

16) Remove the needle, but not the ferrule, from the 10 μL syringe.

17) Remove the small flat retaining nut from the repeating dispenser with the aid of a small coin - a US cent works well. Be careful not to lose the rubber gasket on the syringe side of the retaining nut.

18) With the ratchet arm fully withdrawn insert the syringe without its plunger through the holding ring of the dispenser.

19) Replace the retaining nut, gasket side facing the syringe, and screw it in tightly on the open end of the syringe. CRITICAL STEP. Care should be taken to ensure the syringe is properly centered in the holding ring and that the barrel is aligned parallel to the ratchet arm. Both can be judged by eye. These two requirements are important because if the plunger does not run free and true or it is under strain due to bending or buckling at any point in its travel pressure will build up in the source syringe during loading and catastrophic leakage can occur.

20) Pass the plunger through the gripping ring with the gripper nut unscrewed a few turns and guide the plunger into the open end of the syringe.

21) Depress fully the ratchet arm and move the plunger to the zero graduation mark on the barrel. The syringe is now ready for loading.

22) Move the plunger on one side of the assembled mixing unit (from Step 13) to the zero graduation mark to transfer the homogenous mesophase to the other syringe and the coupler.

23) Disconnect the empty syringe (with its ferrule in place) from the mixing unit and immediately connect the loaded syringe with the coupler attached to the threaded termination of the 10 μL dispensing syringe. CRITICAL STEP. The degree of tightness with which coupling is done is critical, as noted in Step 12. The degree of tightness can be gauged by feel which benefits from practicing the coupling procedure.

24) Load the dispensing syringe by depressing the plunger in the 100 μL syringe so that the mesophase transfers through the coupler. If coupling is tight and the plunger in the dispensing syringe runs free then the latter should begin to move in the direction of motion of the loading plunger as the dispensing syringe fills. CRITICAL STEP. Care should be taken at this point to ensure that the loading plunger is not advanced to such an extent that the dispensing syringe fills to overflowing. This can happen suddenly with the plunger shooting out of the dispensing syringe. If this happens all is not lost because the syringe is loaded and the plunger can simply be reseated. It can then be used in setting up a crystallization trial.

25) Disconnect the loading syringe with the coupler attached from the dispensing syringe.

26) Secure a short, flat-tipped needle to the steel termination of the dispensing syringe and carefully tighten it in place.

27) Clamp the plunger to the ratchet arm by tightening the nut in the gripping ring. CRITICAL STEP. Care should be taken not to over-tighten the nut as this can score and deform the plunger rendering it unusable.

CRITICAL STEP: It is important that the travel range provided to the ratchet arm in the clamped condition be limited to no more than an inch (see Figure 8). Beyond this, the plunger will have a tendency to buckle and delivery can fail.

28) Depress the ratchet drum several times to advance the plunger in the barrel thereby filling the void volume of and loading the needle. This step should be repeated (up to 10 times) until a continuous string of the mesophase emerges from the tip of the needle. The syringe is now ready for use in setting up crystallization trials, which should commence immediately. CRITICAL STEP: It is not advisable to keep the protein-loaded cubic phase for too long before setting up the crystallization trials as some proteins are unstable in the cubic phase without added precipitant.

29) Place the 27-well crystallization plate on a surface raised a few inches above the bench for ease of loading. Optimal contrast and enhanced visibility are achieved when the surface is black, or at least dark. Optionally, the outlet of a small ultrasonic humidifier can be directed toward the crystallization plate to slow evaporation from the dispensed bolus of mesophase during setup. This is especially relevant in areas where ambient relative humidity is low.

30) Homogenize the precipitant solutions, usually in a muti-well block, by shaking, centrifuge them at 3,000 g and room temperature for 3 – 5 minutes (to consolidate solution and to pellet insoluble materials and dust particles), and place next to the crystallization plate for direct and easy access.

31) With the dispensing syringe held vertically in one hand, use the free hand to position the needle tip in the center and directly above the base of well #1. Press the button on the repeating dispenser to expel a bolus of mesophase onto the glass surface. The bolus volume is 200 (100) nL when the standard repeating dispenser is used with a 10 (5) μL syringe. CRITICAL STEP. The tip of the needle should be no more than a few hundred micrometers above the base of the well to ensure proper delivery. If the tip is too far away the mesophase will usually curl up and away from the base; if it is too close the mesophase can ball up around the needle tip to be carried away stuck to the needle. Reproducible delivery is easily achieved. It just takes a little practice.

32) After the three wells in the first row on the plate are loaded with mesophase place 1 μL precipitant solution on top of each mesophase bolus using a 2 μL pipet and standard disposable tips.

33) Repeat Steps 31–32 two more times to fill three rows, each with three wells.

34) As quickly as possible, place a cover slip squarely over the nine filled wells to cover them uniformly. CRITICAL STEP. To effect a water-tight seal use a wooden or a plastic spatula or rod to apply pressure to the cover slip where it makes contact with the exposed sticky surface of the spacer tape. Air gaps can be seen when present between the cover slip and the spacer surface. These can be eliminated by repeatedly working the spatula over the surface of the cover slip with moderate applied pressure. Avoid using anything harder than glass to apply pressure to the cover slip as this may scratch the surface and impair well inspection for crystal growth.

CRITICAL STEP. There are situations where it is necessary to open and reseal wells for purposes of soaking pre-formed crystals with heavy atoms or ligands, for example. In such cases, this is made possible by using 18 mm × 18 mm or 1 inch thick glass coverslips to cover anywhere from one to four wells. A thin layer of vacuum grease is applied to the exposed surface of the spacer ahead of loading. This renders the exposed surface non-sticky but at the same time allows the well to be properly sealed when the coverslip is in place. The coverslip can be removed with relative ease and the well contents adjusted. The well can then be resealed with the original or a new coverslip. The process of opening and closing the well can be repeated as often as needed.

35) Repeat Steps 31–34 until all 27 wells are loaded and sealed. The plate is now ready for inspection and for incubation.

<TROUBLESHOOTING>

Incubation and Tracking Crystallogenesis TIMING 30 – 60 days

36) Place the plates in a temperature-controlled chamber, usually at 20 °C. CRITICAL STEP. It is possible to incubate the plates at other temperatures above and below 20 °C. It is important to note that below about 17 °C the cubic phase of hydrated monoolein is no longer stable and can transform to the lamellar phase (Figure 1) which is not compatible with in meso crystallogenesis. However, the cubic phase exhibits metastability and trials can be set up at lower temperatures but this should be done in the knowledge that the system might convert to the lamellar phase at any time. Shorter chained monoacylglycerols have been designed, synthesized and purified that work at such low temperatures and one has been described in the literature³⁴.

Light-sensitive proteins are usually handled by wrapping the plates in aluminum foil before placing them in the incubation chamber. These can be removed and examined under the microscope in subdued or appropriately colored light.

37) On a regular schedule inspect the wells in crystallization plates using a polarizing light microscope (PLM) with a 10x or 20x objective. The schedule used in the authors’ labs is as follows: day 1, 2, 3, 5, 7, 14, 21, 30, and 60 post-setup. Carefully inspect the mesophase bolus adjusting the depth of focus within the 140 μm-thick sample. Examination should be done both in normal light and between crossed-polarizers.

CRITICAL STEP. Some proteins produce crystals within a few hours; others can take months to grow. It is important to note that crystals can be short-lived and can disappear. It is possible to miss them simply because the plates were not screened often enough. Thus, the frequency with which plates are screened is a judgment call. For those crystals that are slow to grow, the glass sandwich plates offer the advantage that they are essentially water-tight and do not dry out. We have measured this with 96-well plates and found them to undergo no measurable water loss over a period of at least 6 months in a temperature-controlled chamber at 20 °C²⁸.

CRITICAL STEP: The cubic phase is optically isotropic and when sandwiched between two glass plates, as in the case of the crystallization plates used here, optical quality is extremely good. This means that crystals of just a few micrometers in maximum dimension can be seen with ease using a standard light microscope (Figure 9). The presence of crystals can usually be verified by viewing the bolus in the well between crossed polarizers. The cubic phase is not birefringent and appears dark just like the surrounding precipitant solution. In contrast, when non-isotropic crystals are present they usually ‘light up’ and appear as sharp flecks of light in a dark background. Rotating the plate between the crossed polarizers can be used to pick out crystals in different orientations within the bolus.

38) Record the appearance of the mesophase taking care to note whether it is a stiff gel (characteristic of the cubic phase), is smooth and liquid-like (characteristic of the sponge phase) or is completely dissolved. Note too if it is birefringent or optically isotropic. If birefringent, its texture should be noted. Examples of images typically observed are included in Figure 10. Persistent birefringence associated with the lamellar or hexagonal phases suggests that the condition is unlikely to support growth of crystals.

39) Make a note of any crystal-like objects and of their appearance (shape, size, color, orientation, etc.) and number. Carefully examine and note the birefringency of each. Non-birefringent ‘objects’ with sharp edges can be voids in or single domains of the cubic phase (see Figure 11). Initial hits often appear as showers of tiny crystals most obvious when viewed between crossed polarizers. Tryptophan-containing protein crystals can be distinguished from salt, lipid or detergent crystals by imaging with a UV-fluorescence microscope in epi-illumination mode. Excitation and emission wavelengths should range from 280 to 300 nm and 340 to 400 nm, respectively (Korima microscope, Figure 12). Absorption imaging at 280 nm is also possible³⁵ and/or the crystals can be harvested and examined for characteristic X-ray diffraction. Another option is to set up control crystallization trials using protein-free buffer. CRITICAL STEP. In order to carry out UV-fluorescence imaging it is necessary to use either quartz or UV-transmitting glass for the crystallization plate. The 0.5 – 1 mm thick electroverre glass plates from Erie Scientific are suitable for this application. .

40) Carefully examine any precipitated protein, if observed in the mesophase. Birefringent precipitate may provide a useful initial condition for subsequent optimization. It is possible too that on further incubation the protein in the precipitate will convert to a crystal by Ostwald ripening³⁶.

41) If desired, score the drops using a scoring system of the type included in Figure 13 and log the data into a database for subsequent analysis of crystallization trials.

Optimization TIMING 1 – 6 months

42) When a crystallization hit has been recorded, initiate optimization to improve the size and/or diffraction quality. Strategies and protocols for performing optimization that are suitable for use in the in meso method have been described³⁷. Also refer to Box 5 for further information.

Preparing to harvest crystals: Opening the glass cover slip on a crystallization well in a glass sandwich plate. TIMING 5 – 10 min

43) If required, set up an ultrasonic humidifier near the microscope to increase local humidity and to minimize evaporation from the open drop during harvesting.

44) Place the plate on a bench and use the glass-cutting stone to score four lines in the form of a square into the cover slip. The square should be included within the dimensions of the crystallization well and be centered on the mesophase bolus.

CRITICAL STEP. None of the square should extend beyond the well onto the spacer. If it does, the cover slip cannot be removed intact because part of it will remain stuck to the spacer.

<CRITICAL STEP> The glass sandwich crystallization plates are extremely useful for screening conditions that support crystallization. However, they are not particularly easy to harvest from. It is possible to set up in meso crystallization trials in plates from which harvesting is more facile but the conditions for crystallization may not be the same and the new plates will usually require additional screening and optimization. In some cases it is simplest to harvest directly from the glass plates. Harvesting from commercial plates is usually no different to harvesting from glass sandwich plates, as described in steps 52–55. Using plates made of low-birefringency plastic will greatly facilitate the process.

45) Place the scored well into viewing position on the stage of a stereo microscope. Use low magnification, 20 × for example, so that the whole well is in view.

46) With the tip of the tweezers, press on the glass outside and to one side of the square to fracture fully along the score line in the cover slip. Repeat this process along the other three sides.

47) Break holes in the glass cover slip at two opposite corners, just outside the scored perimeter, using the fine tip tweezers.

48) Using a pipette, inject 2 – 3 μL precipitant solution through one of the holes in the cover slip into the crystallization well to saturate the system with precipitant and to reduce the consequences of any dehydration that might occur.

49) Use the angled probe to free the square piece of glass from the rest of the coverslip.

50) Lift the freed glass square with the angled probe away from the well.

CRITICAL STEP. Carefully note the location of the mesophase which can move whilst lifting the square cover. In most cases the mesophase remains on the bottom of the well. Sometimes however it sticks to the upper glass coverslip. In this case rotate the freed square of glass and position it so that it rests on the plate next to the open well with its well side exposed. The angled probe can be used for this procedure.

<TROUBLESHOOTING>

51) IMMEDIATELY, add ~5 μL precipitant solution to the exposed mesophase in the open well and/or on the upturned cover slip.

Harvesting Crystals

TIMING 5 – 20 min.

52) Increase microscope magnification to 60 – 100 x. Rotate the polarizer so that the crystal appears as a bright object on a darker background. Be careful to allow for sufficient light transmittance (in other words do not fully cross the polarizing lenses) so that the loop remains visible whilst the mesophase and the crystal are manipulated.

<TROUBLESHOOTING>

53) Use a Micromount of appropriate size to fish out a crystal.

CRITICAL STEP. Try to take as little mesophase with the crystal as possible. When the crystal resides deep inside the bolus, remove the overlaying mesophase to expose the crystal and to bring it to the surface using one Micromount and then harvest the crystal using a second, clean Micromount. A magnetic wand is used to facilitate handling the Micromount on its magnetic base.

CRITICAL STEP. It is also possible to free a crystal from the mesophase for harvesting by using enzymatic lipid hydrolysis³⁸, detergent solutions³⁹ or mineral oils. However, these assorted manipulations can damage the crystal and lead to poor diffraction quality. In our hands, harvesting directly from the mesophase proved most successful with sensitive crystals and is recommended. Every effort should be made to minimize the amount of mesophase carried over with the crystal to reduce background scatter during diffraction data collection and to facilitate effective cryo-cooling.

54) Harvest the crystal and IMMEDIATELY and RAPIDLY plunge it into clean (water-free) liquid nitrogen (LN2). Allow the Micromount, base and wand tip to cool to LN2 temperature and then transfer the cryo-cooled sample into a pre-cooled vial or a storage puck for storage and shipping.

CRITICAL STEP. The mesophase surrounding the crystal typically preserves the crystal during cryo-cooling. However, in rare cases an additional cryoprotectant may be required. Considerations for the choice of the cryoprotectant are similar to those that apply to the cryo-cooling of soluble protein crystals and for membrane protein crystals prepared using more traditional in surfo methods⁴⁰. Typically glycerol, ethylene glycol, low molecular weight polymers, some organics and light carbonated mineral oils work well.

55) Record in a laboratory notebook the crystal identity, loop type and label, puck number, and the location of the crystal in the puck. Add comments regarding crystal size, shape, color, location and orientation in the loop, and other useful information such as the amount of mesophase carried over with the crystal.

TIMING SUMMARY

Having assembled all the necessary tools, supplies and materials it is possible to prepare the protein/lipid mesophase and to set up a single 27-well crystallization plate in about 30 minutes. With enough mesophase at hand, a second plate can be set up in another 10 minutes. These times apply to the dextrous novice and the experienced worker alike. Actual crystal growing trials can extend from as short a period as a day all the way to months. Once usable crystals (≥25 μm in maximum dimension) are identified harvesting and cryo-cooling can be done in 5 to 20 minutes. However, with some frequency we encounter difficult samples where harvesting a single useful crystal can take an hour. And this was in the hands of an experienced worker. Usually what happens is that experience gained with difficult samples translates into substantially reduced harvesting times as the researcher becomes familiar with the rheology of the mesophase and with the response of the bolus and the crystal to added precipitant, probing tools and harvesting loops.

Optimization based on initial hits can produce crystals of diffraction quality within a single round of optimization that can take as little as a week. More usually however, a series of optimizations is needed where aspects of the protein, the lipid, the precipitant, additives, bolus size, temperature and time are varied systematically. These rounds of optimization can take months to complete, alas with no guarantee of success.

Establishing that a given ‘hit’ arises from the protein, as opposed to some other ingredient in the crystallization well, can be done simultaneously with imaging or viewing with a microscope if the protein is colored. If the protein contains tryptophan and a UV absorption or fluorescence microscope is available, the evaluation can be done in matters of minutes. Longer times are needed to run protein-free controls and to make definitive diffraction measurements.

In preparation for setting up crystallization trials glass sandwich plates must be silanized, perforated spacers applied, and precipitant solutions chosen or formulated as needed. It is preferred to silanize plates and to apply spacers the morning or the day before a trial is to be set up. The time required depends on the number of plates involved. If pre-formulated precipitant solutions are to be used it is simply a matter of selection. At the stage of optimization, precipitant solutions are usually formulated by hand, using a multichannel pipette. The time required depends on the number of conditions to be screened and the number of ingredients in each, tipically ~1 hour per a set of 96 conditions.

When trials are to be performed with mixtures of lipids the corresponding samples should be prepared the previous day. This provides for overnight drying under high vacuum to ensure the lipid is solvent-free at point of use.

Mesophase, lipid and protein characterization, as part of quality control and gaining familiarity with the system under investigation, is desirable but not essential to performing crystallogenesis trials. Typically, such measurements are performed in parallel with ongoing trials. With all of the instrumentation and materials on site it should be possible to make all of the TLC, PLM and SAXS measurements in a single working day. Absorption, fluorescence and CD spectroscopies can also be completed in a day by an experienced researcher with the proper equipment and materials at hand.

Step 13. Problem: Cloudy mesophase

Possible reason 1: Excess protein solution.

Solution: Occasionally an excess of protein solution is used in which case a two-phase system results (see Figure 1). This will give rise to a cloudy dispersion that will not clarify regardless of how much mixing is used. Whilst it is desirable to start out with a one-phase system in the interests of reproducibility, a slight excess of protein solution and a cloudy dispersion should not mean that the crystallization trial is to be abandoned especially when working with a valuable protein and/or lipid. If a clear mesophase is required in such a situation it can be obtained by supplementing the mixture with a small amount (typically < 10 % of the total mixture) of additional lipid. To this end, the homogenized sample should be moved to one of the syringes, the empty syringe disconnected from the coupler and pure lipid added to it. The system is then reassembled, taking care to avoid trapping air bubbles. Upon mixing the overall composition should be such as to restore the pure cubic phase which is optically transparent.

Possible reason 2: Excess lipid.

Solution: Cloudiness in the sample can also originate from inaccurate weighing of the lipid and/or protein solution, or from leaks in the syringe. The consequence here is that the system is less hydrated than is required for cubic phase stability. If this causes the lamellar phase to form or for the lamellar and cubic phases to co-exist (see Figure 1), cloudiness can result. A check on this can be made by transferring all of the homogenized mesophase from one of the syringes and recording total mesophase volume in the other. The value should match closely the sum of the starting lipid and protein solution volumes taking account of the coupler dead volume, which is typically 1.5 μL³³. If the recorded volume is considerably less than expected and the dispersion is cloudy it is important to check the system for leaks and to repeat the mixing protocol with non-leaking syringes.

Possible reason 3: Trapped air.

Solution: Trapping air and bubbles in the lipid and/or protein solution and at the stage where the syringes are combined with the coupler will also give rise to cloudiness and should be avoided if at all possible. Again, this is done in the interests of uniformity and reproducibility.

Possible reason 4: High concentration of detergent in the protein solution.

Solution: A high concentration of detergent in the protein solution can destabilize the cubic phase and transform it fully or partially into a lamellar phase³². This is usually accompanied by the formation of a cloudy dispersion that does not clear with mixing. The problem can be diagnosed by examining the phase texture of a few hundred nanoliters of the dispersion by polarized light microscopy (see Box 2). If the lamellar phase is confirmed the protein can be treated to reduce detergent carry-over. This might involve reducing the detergent concentration in the buffer used to elute the protein from a column by washing with a lower detergent concentration buffer at the concentration step, by using higher molecular weight cut off filters during concentration, and/or by using detergent-binding materials such as Bio-Beads SM-2 (Bio-Rad, Cat. No. 152-3920). It is still possible to perform successful in meso crystallization trials starting from predominately lamellar phase because certain precipitants, highly concentrated salt solutions in particular, can cause the system to revert back to the cubic phase (see ref. 32, for an example of this).

Step 13. Problem: Leakage at the coupler

Possible reason: Loose connection.

Solution: Torque should be maintained during the mixing process by applying a small but steady clockwise, rotating force to the two syringes on either side of the coupler. This is to minimize the likelihood of leakage. We have found that leaks are less of a problem when a tight wrap of Parafilm is applied to the coupler that extends up the base of the two syringes. The ‘bandaging’ is done within about 20 or so strokes when mixing is known to be progressing properly.

Step 13. Problem: Syringe leakage

Possible reason 1: Broken syringe.

Solution: Excessive force applied to the plunger in the barrel of a Hamilton syringe can cause the glass barrel to break or to leak at the junction between the barrel and the threaded steel barrel termination. Syringes that have failed in this way should be discarded.

Possible reason 2: Worn out Teflon plunger tip.

Solution: The Teflon tip at the top of the plunger can wear out from overuse and may start leaking. Replacement plungers for gas-tight syringes are available through Hamilton and are interchangeable between syringes.

Step 13. Problem: Difficulty with mixing

Possible reason: Non-uniform mixing.

Solution: When the lipid and the protein solution come into contact at the beginning of the mixing cycle extra force may be needed to move the contents of the syringe through the coupler. This is to be expected as different phases form depending on the degree of hydration (see Figure 1). However, excessive force should not be used as this will cause the glass barrel to break or the syringe to leak (see previous TROUBLESHOOTING items). Instead it is recommended that the mixing direction be reversed and that less than complete strokes be used to induce local homogenization. With time, the full mixing stroke can be implemented as the syringe contents become more uniform.

Step 35. Problem: Failure to dispense

Possible reason: Clogged needle, coupler.

Solution: It is important not to leave syringes loaded with mesophase open to the atmosphere for more than a few minutes. Otherwise the mesophase can dry out and plug the exposed orifice of a needle, barrel termination or coupler. While these can be cleaned and recovered for reuse it usually involves considerable effort and time. Accordingly, it is best to use the material as quickly as possible and then to dismantle the device and to thoroughly wash the syringes, needles, couplers, ferrules, etc., with methanol followed by milli-Q water and then to dry everything in a stream of compressed dry air or nitrogen gas.

Step 50. Problem: Non-viscous mesophase

Possible reason: Formation of the less viscous sponge phase.

Solution: The sponge phase is a variant of the cubic phase that forms in the presence of certain precipitants²². Depending on the protein harvesting small crystals from the sponge phase can be more difficult than from the cubic phase. The former is much less viscous and can flow. Thus, when the glass coverslip square is broken to access the well capillarity draws the sponge phase and its crystal cargo to the edge of the square where it typically mixes with shards of glass. While the shards are not birefringent and can usually be distinguished from protein crystals their presence complicates harvesting. Further, the sponge phase is ‘stringy’ and excess sponge phase can get carried along with the crystal on the Micromount during harvesting. To overcome these problems a small amount of modified precipitant solution is added to the well as soon as it is opened to regenerate the original, viscous cubic phase. This is achieved by using a lower concentration of sponge-inducing agent in the precipitant solution than was used during the original trial. The conversion takes several minutes at which point harvesting from the cubic phase can proceed as described above. The one risk associated with this solution is that the transformation back to the cubic phase can destroy the crystal which formed under conditions that gave rise to the sponge phase. But in our hands, and especially when working with crystals of GPCR, this approach generally increases the yield of successfully harvested crystals compared to working with the sponge phase directly.

Step 52. Problem: Crystals are not visible under cross-polarized light

Possible reason: Isotropic space group, orientation issue.

Solution: The ability of the crystal to rotate polarized light, and thus its birefringence and visibility, depends in part on the crystal space group and on the orientation of the crystal with respect to the polarizing filters. Sometimes the crystallization plate or the crystal itself must be rotated in the light beam of the microscope to increase or restore crystal birefringency.

Box 2. Problem: Dehydration

Possible reason: Sample exposed to atmosphere for too long.

Solution 1: If the mesophase dries out, even slightly (see phase diagram, Figure 1), it can transform to another phase. The chances of this happening can be reduced by performing Step ii of Section B Polarized Light Microscopy in Box 2 as fast as possible and in a humid atmosphere. Use of a humidifier close to where the work is being done is recommended if problems with drying are encountered. It is also possible to raise local humidity by placing water-saturated tissues in a circle around the microscope slide. Alternatively, the mesophase can be covered with bathing solution as soon as it is placed on the slide and then capped with a cover slip as above. In this case, the bathing solution, which is isotropic, will appear dark when viewed between crossed polarizers. The choice of bathing solution will depend on the composition of the original sample. If it was prepared with protein solution the corresponding protein-free and detergent-free buffer is appropriate. This can be verified by monitoring the PLM texture of the mesophase bolus which should not change with time.

Solution 2: One must also be careful that the system does not lose an excessive amount of water in which case it can transform to the fluid isotropic liquid. The latter is non-birefringent just like the cubic phase and if it is accessed can lead to a case of mistaken phase identity. However, a fluid isotropic bolus can be distinguished by its edges that are uniform and usually circular reflecting the fact that it is a liquid. In contrast, a cubic phase bolus is stiff with edges that appear angular and retain a defined shape.

Box 3. Problem: High UV absorbance

Possible reason: Presence of trace impurities in the lipid.

Solution: Despite the fact that the lipid used can be >99 % pure, as judged by thin layer chromatography and NMR, the UV absorbance of the mesophase can be considerable ⁴¹. This problem of ‘background’ absorbance is exacerbated because the cubic phase sample is close to 60 %(wt/wt) or 1 –2 M lipid. Thus, a small amount of a highly absorbing contaminant in the lipid will contribute significantly to the background signal. Effort has been made to find commercial and homemade lipid with lower absorbance, to no avail. The precise origin of the absorbance signal is not known. Thus, to quantify the spectral properties of the reconstituted protein, all spectra are recorded against an air reference. The analyte protein spectrum is obtained by subtracting the lipid/buffer background from the protein/lipid/buffer sample spectrum.

The Vitamin B₁₂ Transporter, BtuB. A Well-behaved and Robust Membrane Protein

BtuB is an outer membrane protein from the TonB-dependent transporter family responsible for the uptake of cyanocobalamin (Vitamin B₁₂) in gram-negative bacteria. The protein was overexpressed in E. coli and purified as described⁴² using 1.5 %(w/v) octylglucoside and 0.1 %(w/v) LDAO during extraction and purification, respectively. Purified protein was concentrated to 10–15 mg/mL in 10 mM Tris pH 8.0, 0.1 M NaCl, and 0.1 %(w/v) LDAO. Mixing the protein solution with monoolein in a 2/3 weight ratio yielded the cubic-Pn3m phase as evidenced by SAXS (Figure 14; see Figure 15 for representative SAXS patterns). Protein properties were characterized by spectroscopic techniques using microcuvettes and accessories (Figure 16) as described in Box 3. The protein was inserted into the bilayer of the cubic phase which was verified using tryptophan fluorescence quenching by a brominated monoacylglycerol (Figure 17). The secondary structure of the reconstituted protein was undistinguishable from that of the protein in detergent solution as judged by circular dichroism measurements (Figure 17). Finally, the ability of the in meso reconstituted protein to bind Vitamin B₁₂ with native-like affinity was demonstrated by in situ binding measurements²³ (Figure 17).

In meso crystallization was performed with monoolein as the host lipid. The lipidic mesophase laden with protein was dispensed into home-made 96-well glass sandwich crystallization plates using a home-built in meso crystallization robot. Trials were performed with 50 nL protein/lipid dispersion and 1 μL precipitant solution. Plates were incubated at 20 oC for up to 30 days. Crystallization was monitored using an automatic imager RockImager RI24 and manually by light microscope. Initial screening was performed using six 96-well blocks filled with precipitant solution from commercial crystallization kits (Crystal Screen HT, Index HT, SaltRx and MembFac from Hampton Research; Wizard I&II from Emerald BioSystems; MemStart from Molecular Dimensions and JBScreen Membrane from Jena Bioscience). Small crystals of the type shown in Figure 18 were found to form in the presence of MPD, pentaerythritol propoxylate (5/4 PO/OH), PEG 550 MME, jeffamine M-600, 1,4-butanediol and potassium thiocyanate. Conditions containing MPD were optimized using coarse concentration-pH grid screens ([MPD] range 6 – 14 %(v/v) with 2 %(v/v) steps, pH range 4 – 8 with 1 pH unit steps), followed by screening with 48 different salts as additives (two concentrations for each salt in the 0.1 – 0.5 M range) and finishing with fine concentration-pH grid screens ([MPD] range 8 – 12 %(v/v) in steps of 1 %(v/v), salt concentration range 0.05 – 0.3 M in 0.05 M steps, pH range 6 – 7 in pH unit steps of 0.5). About five subsequent rounds of optimization (using visual, not diffraction quality cues) were needed to obtain ~100 μm-sized BtuB crystals. The best crystals were grown in 10–12 %(v/v) MPD, 0.1–0.2 M ammonium formate, and 0.1 M MES, pH 6.5. Crystals of BtuB for crystallographic measurement were grown in a 72-well Nunc microbatch plate using a fine concentration-pH grid screen around the best conditions found with sandwich plates. 200 nL lipid/protein dispersion was used in each well in conjunction with 3 μL precipitant solution. Wells were sealed with transparent tape. Crystals grew to their full size (~100 × 60 × 25 μm³) in 10–14 days (Figure 9D). Harvesting was done after 14 days using Hampton cryo-loops (0.1 mm diameter) and crystals were cryocooled and stored in liquid nitrogen. Lipidic cubic phase that had swollen in the presence of MPD and that had transformed into a sponge phase had the consistency of a viscous liquid, which did not interfere with harvesting. MPD and lipid served as cryoprotectants and no additional cryoprotectant was used.

Diffraction data from several crystals were collected on the F1 beamline at the Cornell High-Energy Synchrotron Source (CHESS), Ithaca, NY. Best sets from two different crystals were merged to obtain complete data at 1.95 A.

Light Harvesting Complex II, LH2. Working with a Chromophore-Containing Protein and Short-lived Crystals

Light-harvesting complex 2 (LH2) functions as an antenna to trap light energy and to pass it to the reaction center in photosynthetic bacteria. Wild type LH2 was purified using 0.05 %(w/v) LDAO from Rhodopseudomonas acidophila as described⁴³ and was concentrated to 20 – 30 mg/mL in 20 mM Tris pH 8.0, 0.05 %(w/v) LDAO.

Samples for use in crystallization trials were prepared with 60 %(w/w) monoolein and 40 %(w/w) protein solution (20–30 mg protein/mL) using the lipid mixing device. The UV-visible absorption spectrum of LH2 reconstituted in the cubic phase was similar to that obtained with the protein in detergent solution with one difference; a slight distortion in the bacteriochlorophyll absorption bands was observed and attributed to constraints imposed on the protein by the bilayer curvature of the mesophase (Figure 19). Initial crystallization screening was performed with the in meso robot. To this end, each well in a home-made, 96-well glass sandwich crystallization plate was loaded with 50 nL lipid/protein dispersion followed by 1 μL precipitant solution at room temperature (20–22 °C). Four commercial screens were performed in duplicate. Incubation was carried out in a temperature-regulated dark chamber at (20 °C). Crystallization progress was evaluated several times in a 2-week period using an automatic imager RockImager RI54 and manually by microscope. Initial crystal hits were found in a number of conditions containing precipitants that cause the cubic phase to swell, such as MPD, Pentaerythritol Propoxylate (PPO), 1,4 - butanediol, and Jeffamine M-600. Optimized conditions leading to diffraction quality crystals included 20 %(v/v) PPO, 0.1 M potassium acetate, and 0.1 M Tris-HCl pH 8.0²². Crystals typically grew as long rods (~200 × 30 × 10 μm³) for 3–7 days (Figure 9B). After 7–10 days the crystals began to degrade. Therefore harvesting was carried out within a week of set up to ensure best crystal quality.

A full 2.45 Å dataset was collected from a single crystal on the F1 beamline at CHESS.

Adhesin, OpcA. Use of Inclusion Bodies

OpcA is an integral outer membrane adhesin protein from Neisseria meningitidis, the causative agent of meningococcal meningitis and septicemia. It binds to sialic acid (SA)-containing polysaccharides on the surface of epithelial cells. Recombinant OpcA was obtained by expression of the protein into inclusion bodies, followed by solubilization in guanidine HCl and rapid dilution into buffer containing LDAO⁴⁴. Subsequent purification was carried out as described previously ⁴⁴. After the last purification step the protein was precipitated in ethanol and stored in small aliquots at −20 °C. Before crystallization, a vial of the protein-in-ethanol suspension was centrifuged for 15 min at 14,000 g. Ethanol was removed and the pellet was dissolved in 0.1 %(w/v) DDM, 50 mM bis-tris propane pH 7.0 to 0.2 mg protein/mL. The protein was then concentrated using a Microcon spin concentrator with 30 kDa cut off to 10–15 mg/mL.

In meso crystallization was performed with monoolein as the host lipid. The lipidic mesophase laden with protein was prepared as a 3/2 by volume mix of lipid and protein solution and dispensed into home-made 96-well crystallization plates using a home-built in meso crystallization robot. Trials were performed with 50 nL protein/lipid dispersion and 1 μL precipitant solution. Plates were incubated at 20 oC for up to 30 days. Crystallization was monitored using an automatic imager RockImager RI54 and manually by light microscope. Initial screening was performed using six 96-well blocks filled with precipitant solution from commercial crystallization kits (Crystal Screen HT, Index HT, SaltRx and MembFac from Hampton Research; Wizard I&II from Emerald BioSystems; MemStart from Molecular Dimensions and JBScreen Membrane from Jena Bioscience). Small crystals were found to form in the presence of PEG 400 and lithium sulfate. Initial conditions were optimized using concentration-pH grid screens and different salts as additives to obtain ~70 μm-sized crystals (Figure 9C). The best crystals were grown in 18 %(v/v) PEG 400, 0.1 M potassium sulfate, 0.05 M Hepes, pH 7.0²⁴.

Crystals of apo-OpcA for crystallographic measurement were grown in a 96-well Greiner sitting drop plate. 200 nL lipid/protein dispersion was used in each well in conjunction with 2 μL precipitant solution. Each reservoir contained 50 μL of the corresponding precipitant solution. Wells were sealed with transparent tape. Crystals grew to their full size (25 × 25 × 70 μm³) in 20 – 30 days. Harvesting was done after 24 days using Hampton cryo-loops (0.05 – 0.1 mm diameter) and crystals were cryocooled and stored in liquid nitrogen. Crystals were harvested directly from the lipidic cubic phase and were frozen immediately without further treatment. During harvesting care was taken to pick up as little mesophase as possible. PEG 400 and lipid served as cryoprotectants and no additional cryoprotectant was used.

A full dataset at 1.95 Å was collected from a single crystal on the F1 beamline at CHESS.

Engineered human β₂-Adrenergic Receptor, β₂AR-T4L. Working with a Highly Unstable Protein, Micro-crystals and a Lipid Additive

The β₂-adrenergic receptor (β₂AR) is a member of the G protein-coupled receptor (GPCR) family. It regulates cardiovascular and pulmonary function in response to elevated levels of the endogenous hormones adrenaline and noradrenaline. To increase its stability and aid in crystallization the human β₂AR was modified as follows⁴⁵:

1. The third intracellular loop was replaced with a compact and stable protein lysozyme from T4 phage,

2. The C-terminal 48 amino acids were removed, and

3. A glycosylation site at Asn187 was mutated to Glu.

The engineered protein, called β₂AR-T4L, was expressed in sf9 insect cells infected with baculovirus. The protein was solubilised in DDM and purified as described in ref. 45. Purified protein was treated with iodocetamide to block free sulfhydryls and additionally stabilized by a partial inverse agonist carazolol and concentrated to 30 – 50 mg/mL for crystallization trials. Initial crystallization screening in lipidic cubic phase was performed with glycosylated and de-glycosylated (treated with PNGase F) receptors using monoolein as host lipid against seven commercial 96-well screen kits in duplicate. Trials were set up in 96-well glass sandwich plates using 50 nL of lipidic cubic phase and 0.8 μL of precipitant solution per well dispensed by the in meso robot. Initial hits consisting of showers of micro-needles (< 5 μm in length; Figure 20) were obtained only with de-glycosylated protein at three different conditions, all containing PEG 400, lithium sulphate and a buffer at pH between 6.5 and 8. Precipitant and salt concentration pH optimization grids combined with salt identity optimization provided useful ranges and optimal values for PEG 400 and salt (sodium sulphate was identified by screening for salt identity) concentrations and buffer pH, and yielded slightly larger crystals measuring 15 × 5 × 1 μm³. The most dramatic improvement came from screening with soluble and lipophilic additives. We have found that the addition of cholesterol to the host lipid and of 1,4-butanediol to the precipitant solution significantly improved the shape and the size of crystals to an average of 30 × 10 × 5 μm³. Finally, the conditions were fine-tuned, as a result of which the buffer was changed from Hepes to Bis-tris propane, the lipidic cubic phase bolus size was reduced from 50 to 20 nL, and the protein concentration was increased from 30 to 50 mg/mL. To arrive at the final crystallization conditions consisting in 30–35 %(v/v) PEG 400, 0.1–0.2 M sodium sulfate, 0.1 M Bis-tris propane pH 6.5–7.0 and 5–7 %(v/v) 1,4-butanediol as the precipitant solution and 8–10 %(w/w) cholesterol in monoolein as the host lipid, we formulated thirty five 96-well screens and performed over 15,000 trials. Therefore, robotics were indispensable to achieving success in this project. The fully optimized crystals were grown for 3 – 6 days and reached a maximum size of 40 × 20 × 7 μm³²⁵ (Figure 9E).

Extensive attempts to reproduce crystals in commercial plates for microbatch, and for sitting and hanging drop vapour diffusion resulted in significantly smaller crystals. Therefore crystals were harvested directly from the glass sandwich plates. Crystals were scooped from lipidic cubic or sponge phase using MiteGen 30 or 50 μm MicroMounts and plunged into liquid nitrogen. Due to difficulties of harvesting small crystals from sponge phase (see Troubleshooting to Step 53) the concentration of PEG 400 was lowered to ~26 %(v/v) to transform the sponge phase into a cubic before harvesting. Care was taken to reduce the amount of mesophase that came with the crystal onto the mount so as to minimize unwanted background X-ray scattering. Attempts to dissolve the mesophase either by increasing the concentration of PEG 400 or by using mineral oils typically resulted in a reduction in the diffraction power of the crystals.

Because the crystals were so small it was necessary to use an equally small but highly collimated X-ray beam for diffraction data collection. Accordingly, data were collected using a 10 μm-diameter minibeam on the 23 ID-B beam line (GM/CA CAT) at the Advanced Photon Source, Argonne, IL. Crystals diffracted to better than 2.2 Å resolution but were susceptible to radiation damage. Therefore, a complete, low resolution dataset, collected with a highly attenuated beam, was merged with 31 high-resolution small sets of 10–20 degrees each to obtain complete data at 2.4 A resolution²⁵.

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