Name: abcg5/g8 crystallization in a lipidic bicelle environment for x-ray crystallography
Uploaded: 2023-08-25T14:40:00.000+00:00
Duration: 6 min 47 s
Description: Watch this Scientific Journal Video about A Bicelle Crystallization Setup for ABC Transporter Membrane Proteins to Advance Drug Development at JoVE.com

This work is supported by a Natural Sciences and Engineering Research Council Discovery Grant (RGPIN 2018-04070) and a Canadian Institutes of Health Research Project Grant (PJT-180640) to JYL. This protocol is based on the original reports in ABCG5/G8 crystal structures reported earlier by Farhat et al.22 and Lee et al.23.

1. Cloning and protein expression
<ol>
	<li>Clone the human ABCG5/G8 gene into Pichia pastoris yeast following previous protocols25,26. Briefly, derive pSGP18 and pLIC expression vectors from pPICZB. Add a tag that encodes a rhinovirus 3C protease site followed by a calmodulin binding peptide (CBP) to the C-terminus of ABCG8 cDNA (pSGP18-G8-3C-CBP).
	<ol>
		<li>Add a six-histidine tag separated by a glycine (His6GlyHis6) to the C-terminus of ABCG5 cDNA (pLIC-G5-H12). Co-transform the plasmids into Pichia strain KM71H using electroporation. 
		NOTE: Please see the Table of Materials for the details of the plasmids, media and buffers used.</li>
		<li>Grow transformed yeast cells on MD agar plates at 28 &#176;C.</li>
	</ol>
	</li>
	<li>After 1-2 days, select 10-12 colonies and inoculate them into 10 mL of minimal glycerol yeast nitrogen base (MGY) media using 50 mL centrifuge tubes for small-scale culture.
	<ol>
		<li>Make three small holes in the centrifuge tube lid for better aeration. Allow cells to grow at 28 &#176;C with constant shaking at 250 rpm until the optical density at 600 nm (OD600) reaches 10, usually taking 1-2 overnights. 
		NOTE: Cell growth usually takes between 12-24 h.</li>
	</ol>
	</li>
	<li>On the next day, take 1 L of sterile MGY media and inoculate it with the primary culture in a 2.4 L flask. Incubate the flask at 28 &#176;C in a shaker incubator at 250 rpm for 24 h.
	<ol>
		<li>To maintain pH between 5-6, add 10% ammonium hydroxide (NH4OH) until the pH stabilizes.</li>
	</ol>
	</li>
	<li>Adjust the pH and induce protein expression by adding 1 mL of pure methanol per 1 L culture (0.1 % (v/v) methanol). 
	NOTE: Feed cells subsequently with 5 mL pure methanol per liter culture (0.5 % (v/v) methanol) every 12 h for a total duration of 36-48 h.</li>
	<li>Harvest cells by centrifuging at 15,000 x g for 30 min at 4 &#176;C.</li>
	<li>Collect cell pellets and resuspend them in lysis buffer (0.33 M sucrose, 0.3 M Tris-Cl pH 7.5, 0.1 M aminohexanoic acid, 1 mM EDTA, and 1 mM EGTA) to a concentration of 0.5 g/mL. Store the suspension at -80 &#176;C. Typically, one can recover 30 &#177; 5 g of cell mass from 1 L of cultured cells. 
	&#8203;NOTE: Store cell pellets directly in the freezer or perform immediate resuspension in lysis buffer for membrane preparations.</li>
</ol>
2. Preparation of microsomal membrane
<ol>
	<li>Thaw the cells and add protease inhibitors (2 &#181;g/mL leupeptin, 2 &#181;g/mL pepstatin A, 2 mM PMSF, see Table of Materials).
	<ol>
		<li>To further lyse cells, use an ice-chilled emulsifier or microfluidizer (see Table of Materials) at 25,000-30,000 psi. Repeat this process 3-4 times.</li>
		<li>Centrifuge to remove cell debris: spin at 3,500-4,000 x g for 15 min, followed by a second spin at 15,000 x g for 30 min. Maintain both spins at 4 &#176;C.</li>
	</ol>
	</li>
	<li>To isolate microsomal membrane vesicles, transfer the supernatant to ultracentrifuge tubes and subject it to ultracentrifugation at 2,00,000 x g for 1.5 h at 4 &#176;C.
	<ol>
		<li>Resuspend the membrane pellet in 50 mL of Buffer A (50 mM Tris-Cl pH 8.0, 100 mM NaCl, and 10% glycerol) using a dounce homogenizer. Store the suspension at -80 &#176;C.</li>
	</ol>
	</li>
</ol>
3. Protein preparation-purification of heterodimers
<ol>
	<li>Thaw the frozen microsomal membranes, and adjust the concentration to 4-6 mg/mL using solubilization buffer. The buffer should contain 50 mM Tris-HCl, pH 8.0, 100 mM NaCl, 10% glycerol, 1% (w/v) &#946;-dodecyl maltoside (&#946;-DDM), 0.5% (w/v) cholate, 0.1 % (w/v) cholesteryl hemisuccinate (CHS), 5 mM imidazole, 5 mM &#946;-mercaptoethanol (&#946;-ME), 2 &#181;g/mL leupeptin, 2 &#181;g/mL pepstatin A, and 2 mM PMSF (see Table of Materials). 
	NOTE: One can mix equal volumes of the membrane preparation and the solubilization buffer, or use 2x solubilization buffer without protease inhibitors and reducing agents. Short boiling of the buffer helps dissolve CHS efficiently. For purification, use only the 4 &#176;C-cooled buffer.
	<ol>
		<li>Stir the mixture at medium speed for 1 h at 4 &#176;C. Follow this with additional stirring at room temperature (RT) for 20-30 min.</li>
		<li>Centrifuge the mixture at 1,00,000 x g for 30 min at 4 &#176;C to remove insoluble membranes. Collect the solubilized supernatant and add 20 mM imidazole and 0.1 mM TCEP.</li>
	</ol>
	</li>
	<li>Perform affinity column chromatography26: bind the solubilized supernatant to pre-equilibrated Ni-NTA beads (10-15 mL) (see Table of Materials) in Buffer A (step 2.2.1) overnight. 
	NOTE: Avoid using glycerol in running buffers from this point onward.
	<ol>
		<li>Wash the column twice with 10 column volumes of Buffer B (50 mM HEPES, pH 7.5, 100 mM NaCl, 0.1 % (w/v) &#946;-DDM, 0.05 % (w/v) cholate, 0.01 % (w/v) CHS, 0.1 mM TCEP) containing 25 mM imidazole.</li>
		<li>Wash the column with 10 column volumes of Buffer B containing 50 mM imidazole.</li>
		<li>Elute the protein using Buffer C (Buffer B with 200 mM imidazole).</li>
		<li>Add 1 mM TCEP (see Table of Materials) and 10 mM MgCl2 to the eluted proteins.</li>
		<li>Validate the eluted fractions on an SDS-PAGE gel to confirm the correct protein size26. 
		NOTE: Typical protein yield (1st Ni-NTA): 10-20 mg protein per 6 L culture. Use DDM at 10x or 5x of its critical micelle concentration (CMC), approximately 0.01%. This protocol employs 0.1% DDM.</li>
		<li>Dilute the peak fractions from Ni-NTA elution with an equal volume of Buffer D1 (Buffer B with 1 mM CaCl2, 1 mM MgCl2), mix, and load the protein fractions onto a CBP column (3-5 mL) (see Table of Materials) that has been pre-equilibrated with CBP wash Buffer D1.</li>
		<li>Perform sequential washes on the CBP column to exchange detergents using Buffer D1 and D2 (Buffer B with 1 mM CaCl2, 1 mM MgCl2, 0.1% (w/v) decyl-maltose neopentyl glycol (DMNG), without &#946;-DDM): first, wash with 3 column volumes of D1; second, wash with 3 column volumes of D1:D2 (3:1, v/v); third, wash with 3 column volumes of D1:D2 (1:1, v/v); fourth step, wash with 3 column volumes of D1:D2 (1:3, v/v), followed by 6-10 column volumes of D2.</li>
		<li>Elute the protein using CBP wash Buffer D2 with 300 mM NaCl in 1 mL fractions from the CBP column (total 10 mL). Concentrate the eluted fractions to 1-2 mL. 
		NOTE: The typical protein yield (1st CBP) is 5-15 mg protein per 6 L culture. Maltose neopentyl glycol (MNG) detergents enhance purified protein storage at 4 &#176;C. Both DMNG and Lauryl MNG (LMNG) were used, with DMNG yielding better X-ray diffracting crystals. Use DMNG at 10-20x of its critical micelle concentration (CMC), approximately 0.003%. This protocol used 0.1% DMNG. A fraction of the CBP eluate can be further purified by gel-filtration chromatography (step 4.4.) to analyze proteins&#39; ATPase activity or assess mono-dispersity through transmission electron microscopy (TEM).</li>
	</ol>
	</li>
</ol>
4. Protein preparation-pre-crystallization treatment
<ol>
	<li>Cleave the N-linked glycans and CBP tags using endoglycosidase H (Endo H, ~0.2 mg per 10-15 mg purified protein) and HRV-3C protease (~2 mg per 10-15 mg purified protein), respectively (see Table of Materials). Incubate overnight at 4 &#176;C.</li>
	<li>During the Endo H and 3C protease incubation, perform reductive alkylation on the pooled proteins. Begin by incubating with 20 mM iodoacetamide (see Table of Materials) overnight at 4 &#176;C. Follow this with a 1 h incubation with an additional 2 mM iodoacetamide on ice. 
	NOTE: This step further stabilizes protein storage for up to one month at 4 &#176;C.</li>
	<li>Employ a second CBP column (1-2 mL) to separate the cleaved CBP tag. Use Buffer D2 for this process. 
	NOTE: The typical protein yield (2nd CBP) is 5-10 mg protein per 6 L culture.</li>
	<li>Purify the CBP tag-free protein using gel filtration chromatography. The buffer should contain 10 mM HEPES, pH 7.5, 100 mM NaCl, 0.1% (w/v) DMNG, 0.05 % (w/v) cholate, and 0.01% (w/v) CHS. 
	NOTE: The typical protein yield (gel-filtration) is 2-8 mg protein per 6 L culture. During this step, the absence of a DDM peak (~65 kD) during gel filtration indicates successful detergent exchange to DMNG.</li>
	<li>Modify the pooled protein fractions through reductive methylation: add 20 mM dimethylamine borane (DMAB, see Table of Materials) and 40 mM formaldehyde to the protein. Incubate for 2 h at 4 &#176;C on an oscillatory shaker. Add 10 mM DMAB.
	<ol>
		<li>Repeat step 4.5, including the addition of 10 mM DMAB, and incubate overnight (12-18 h) at 4 &#176;C.</li>
		<li>Halt the reaction by adding 100 mM Tris-Cl, pH 7.5.</li>
	</ol>
	</li>
	<li>Load the methylated protein onto a 2 mL Ni-NTA column pre-equilibrated with 100 mM Tris-Cl, pH 8.0, and 100 mM NaCl.
	<ol>
		<li>Wash the column using 10 column volumes of wash buffer (10 mM HEPES, pH 7.5, 100 mM NaCl, with 0.5 mg/mL DOPC: DOPE (3:1, w/w), 0.1% (w/v) DMNG, 0.05% (w/v) cholate, 0.01% (w/v) CHS).</li>
		<li>Elute the relipidated protein using elution buffer (10 mM HEPES, pH 7.5, 100 mM NaCl, 200 mM imidazole, 0.5 mg/mL DOPC: DOPE (1:1, w/w), 0.1% (w/v) DMNG, 0.05% (w/v) cholate, 0.01% (w/v) CHS). 
		NOTE: The typical protein yield (2nd Ni-NTA) is 1-5 mg protein per 6 L culture.</li>
		<li>Pass the protein eluates through a PD-10 desalting column pre-equilibrated with the buffer used in step 4.4.</li>
	</ol>
	</li>
	<li>Incubate the desalted and relipidated protein overnight with cholesterol (prepared in isopropanol or ethanol) to a final concentration of ~20 &#181;M.
	<ol>
		<li>The next morning, remove the precipitant by ultracentrifugation at 1,50,000 x g for 10 min at 4 &#176;C. Collect the supernatant.</li>
		<li>Concentrate the protein to a final concentration of 30-50 mg/mL using a 100 kDa cutoff centrifugal concentrator.</li>
		<li>Remove the precipitant using a benchtop refrigerated centrifuge at the top speed for 30 min at 4 &#176;C.</li>
		<li>Keep the supernatant on ice at 4 &#176;C and establish crystallization conditions in the bicelles. 
		&#8203;NOTE: The concentrated proteins should be used for crystal growth within a week. Do not freeze the proteins.</li>
	</ol>
	</li>
</ol>
5. Protein crystallization in bicelles
<ol>
	<li>Prepare a 10% bicelle stock solution with DMPC lipids and CHAPSO detergent in a 3:1 (w/w) ratio (see Table of Materials). 
	NOTE: Use CHAPSO at 5x of its critical micelle concentration (CMC), approximately 0.5%. This maintains detergent concentration around its CMC in the protein-bicelle mixture (step 5.2).
	<ol>
		<li>Add deionized H2O-dissolved detergent (CHAPSO) to pre-dried lipids (mixture of 5 mol% cholesterol and 95 mol% DMPC). 
		NOTE: Prepare various lipid compositions in chloroform, and dry them in a glass test tube using a nitrogen gas stream at RT. Eliminate residual solvents by placing them in a vacuum chamber overnight, forming a thin lipid layer.</li>
		<li>Resuspend lipids and detergent using a water-bath sonicator.</li>
		<li>Sonicate the bicelle mixture in ice-chilled water using continuous power until the solution becomes transparent. 
		NOTE: Use hearing protection and ensure a sufficient ice supply to keep the mixture in a liquid phase.</li>
		<li>Remove undissolved components with a 0.2 &#181;m centrifugal filter (see Table of Materials). 
		NOTE: Store the aliquoted bicelle solution at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Create a protein/bicelle mixture on ice by gently combining 10% bicelles (step 5.1.4) and proteins (step 4.7.4) in a 1:4 (v/v) ratio, achieving a final protein concentration between 5-10 mg/mL.</li>
	<li>Incubate the protein and bicelle mixture on ice for 30 min.</li>
	<li>Set up crystallization conditions in a hanging drop vapor diffusion format using 48-well plates.
	<ol>
		<li>Mix equal volumes (0.5 or 1 &#181;L) of protein/bicelle mixture and crystallization reservoir solution containing 1.6 M-2.0 M ammonium sulfate, 100 mM MES (pH 6.5), 0%-4% PEG 400, and 1 mM TCEP (see Table of Materials). 
		NOTE: Create a matrix of the reservoir solution prior to each experiment, adjusting ammonium sulfate (1.6-2.0 M) and PEG 400 (0%-4%).</li>
		<li>Incubate for crystallization at 20 &#176;C.</li>
		<li>Check the crystallization trays the next day to ensure proper cover glass sealing.</li>
		<li>Monitor crystal growth at least once daily. High-quality crystals usually appear within 1-2 weeks, measuring 50-150 &#181;m x 20-50 &#181;m x 2-5 &#181;m. 
		NOTE: Crystals may take longer to form at lower protein concentrations. Mature crystals should be harvested within one month.</li>
		<li>Soak protein crystals in 0.2 M sodium malonate and flash freeze them in liquid nitrogen using 50 or 100 &#181;m cryo-loops. 
		NOTE: If an X-ray diffractometer is available, test a few crystals with a 15-30 min X-ray beam exposure to reveal diffraction up to 5 &#197;. Higher-resolution diffraction requires a synchrotron light source. Using 0.2 M sodium malonate as a cryo-protectant, a crystal measuring 100 &#181;m x 50 &#181;m x 2 &#181;m can provide around 90 diffraction image frames with synchrotron X-ray.</li>
	</ol>
	</li>
</ol>

Crystallization of ABC Transporter Membrane Proteins in Lipidic Bicelles

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Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystal+structure+of+the+human+sterol+transporter+ABCG5/ABCG8.">Crystal structure of the human sterol transporter ABCG5/ABCG8.</a> Nature. 533 (7604), 561-564 (2016).</li><li>Ujwal, R., Bowie, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystallizing+membrane+proteins+using+lipidic+bicelles.">Crystallizing membrane proteins using lipidic bicelles.</a> Methods. 55 (4), 337-341 (2011).</li><li>Johnson, H., Lee, J. Y., Pickert, A., Urbatsch, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bile+acids+stimulate+ATP+hydrolysis+in+the+purified+cholesterol+transporter+ABCG5/G8.">Bile acids stimulate ATP hydrolysis in the purified cholesterol transporter ABCG5/G8.</a> Biochemistry. 49 (16), 3403-3411 (2010).</li><li>Wang, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Purification+and+ATP+hydrolysis+of+the+putative+cholesterol+transporters+ABCG5+and+ABCG8.">Purification and ATP hydrolysis of the putative cholesterol transporters ABCG5 and ABCG8.</a> Biochemistry. 45 (32), 9929-9939 (2006).</li><li>Faham, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystallization+of+bacteriorhodopsin+from+bicelle+formulations+at+room+temperature.">Crystallization of bacteriorhodopsin from bicelle formulations at room temperature.</a> Protein Science. 14 (3), 836-840 (2005).</li></ol>

The authors do not have anything to disclose.

The challenges associated with crystallizing membrane proteins have prompted the development of lipid-bilayer-driven crystallization methods, such as the bicelle27 or lipid cubic phase (LCP)14 approaches. However, achieving successful crystallization of membrane proteins still hinges on the critical and sometimes bottlenecked step of protein preparation. Notably, ABC transporters present a formidable hurdle in growing crystals suitable for X-ray crystallography. This protocol provides comprehensive hands-on guidance for streamlining the preparation of human ABCG5/G8 sterol transporter and fostering crystal growth through the bicelle crystallization approach.
A key consideration in devising this protocol was the imperative for a substantial protein yield in the initial phases of protein purification, allowing for a certain degree of protein loss during pre-crystallization treatment (Figure 3). Common strategies for addressing this challenge involve extensive protein engineering, utilization of diverse expression hosts, and exploration of orthologs or homologs, among other approaches. Nevertheless, with this seemingly intricate procedure, a number of pivotal steps have been identified that underpin the protocol's success and also provide insights into potential limitations that may arise when studying other ABC transporters or membrane proteins in general.
Firstly, this protocol employs thorough centrifugation at each step to minimize protein aggregation. Additionally, continuous monitoring of the thermostability of the purified proteins is crucial. Electron microscopy is utilized to verify protein monodispersity, while analytical gel filtration tracks protein stability over time (Figure 2). Alternative techniques like circular dichroism (CD) or differential scanning calorimetry (DSC) could also be incorporated. Furthermore, the incorporation of lipids at specific stages is essential to maximize both the activity and crystallogenesis of the purified ABCG5/G8. For instance, cholate and CHS are necessary to exhibit measurable ATP hydrolysis; phospholipids are indispensable for maintaining the stability of methylated proteins; and cholesterol is a requisite component of the bicelle solution, fostering crystal growth suitable for high-resolution X-ray diffraction (Figure 4).
In essence, the entire procedure can be accomplished within a week's worth of effort. In contrast to LCP, the retrieval of crystals from hanging-drop crystallization trays is straightforward. Looking ahead, with a substantial protein yield (approximately 10 mg), this protocol is readily adaptable for developing crystallographic investigations involving ABCG5/G8 mutants or other transporter proteins. This is particularly pertinent for cases that currently evade visualization through electron microscopy.

ATP-binding cassette (ABC) transporters constitute a superfamily of membrane proteins responsible for diverse ATP-dependent transport processes across biological membranes1,2,3,4,5. These transporter proteins are implicated in cardiovascular diseases and play a significant role in facilitating cholesterol efflux to the bile for subsequent excretion in the liver. Consequently, cholesterol metabolism and balance have garnered considerable interest over the years6. A specific mechanism involved in the elimination of cholesterol and other sterols from the body involves members of the human ABCG subfamily, notably the heterodimeric ABCG5/G87,8,9,10. Mutations in either of these genes disrupt the heterodimer, leading to loss of function and causing sitosterolemia, a disorder affecting sterol trafficking11,12,13. Given the disease&#39;s relevance and their role in promoting cholesterol efflux, sterol transporters have attracted significant attention. Nevertheless, the intricate details of their molecular mechanism and substrate selectivity remain largely undisclosed. Thus, the elucidation of the crystal structure of ABCG5/G8 is a crucial stride toward comprehending the mechanisms and downstream functions in cholesterol transport.
Membrane proteins require anchoring within membranes to fold and function correctly. Consequently, extracting membrane proteins from their natural environment often results in protein instability, misfolding, and loss of function14,15. These challenges underscore the primary hurdles faced in membrane protein crystallization. However, the reconstitution of proteins into synthetic detergent bilayers, like bicelles, has emerged as a solution to this predicament, enabling the maintenance of membrane proteins within a native-like bilayer milieu16. Bicelles are assemblies of synthetic phospholipids and detergents suspended and solubilized in water. Notably, they adopt a bilayer structure that mimics biological membranes16,17,18. Bicelles can transition between liquid and gel phases based on temperature and viscosity. Bicelle crystallization capitalizes on the small bilayer discs and low viscosity at reduced temperatures, facilitating thorough mixing of proteins and bicelle solutions. The size of the bicelles depends on the detergent-to-lipid ratio during preparation19,20. The prevalent detergents for bicelle formation include 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxy-1-propanesulfonate (CHAPSO), along with 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS) and 1,2-ditridecanoyl-sn-glycerol-3-phosphocholine (DHPC)21. These detergents are used in conjunction with lipids such as di-myristoyl-phosphatidylcholine (DMPC) and 1-palmitoyl-2-oleoyl-phosphatidylcholine (POPC). Furthermore, recent studies have demonstrated the full functionality of membrane proteins within bicelles under physiological conditions. For example, Lee and colleagues successfully crystallized and reported the crystal structure of ABCG5/ABCG8 based on a lipid bilayer22,23. In the crystallization process, protein-bicelle mixtures can be accommodated using standard equipment, including high-throughput crystallization robots24. The feasibility of utilizing bicelles, however, hinges on the proteins&#39; thermostability due to the crystallization conditions at higher temperatures. Nevertheless, when compared to other techniques, the requisite crystallization conditions for membrane proteins generally remain mild, involving low concentrations of precipitant, salt, and buffer. This renders both protein-bicelle mixtures and vapor diffusion effective and easily implementable tools for structural studies of membrane proteins.
This protocol outlines essential steps in protein preparation and bicelle crystallization for determining the X-ray crystal structure of ABCG5/G8 at high resolution (Figure 1).

<table><tbody><tr><td>ABCG5</td><td>National Institute of Health collection&amp;nbsp;</td><td></td><td>NCBI accession number NM_022436</td></tr><tr><td>ABCG8</td><td>National Institute of Health collection&amp;nbsp;</td><td></td><td>NCBI accession number NM_022437</td></tr><tr><td>&amp;Auml;KTA FPLC system&amp;nbsp;</td><td>Cytiva (formerly GE Healthcare Life Sciences)</td><td></td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Wisent</td><td>600-024-CG</td><td>Anhydrous</td></tr><tr><td>CBP</td><td>Agilent</td><td>214303</td><td>Calmodulin binding peptide affinity resin</td></tr><tr><td>Centrifugal concentrators (Vivaspin)</td><td>Sartorius</td><td></td><td></td></tr><tr><td>CHAPSO&amp;nbsp;</td><td>Anatrace</td><td>C317</td><td>Anagrade</td></tr><tr><td>Cholesterol</td><td>Anatrace</td><td>CH200</td><td></td></tr><tr><td>CHS</td><td>Steraloids</td><td>C6823-000</td><td></td></tr><tr><td>DMAB</td><td>MilliporeSigma</td><td>180238</td><td>97%</td></tr><tr><td>DMNG</td><td>Anatrace</td><td>NG322</td><td></td></tr><tr><td>DMPC</td><td>Anatrace</td><td>D514</td><td></td></tr><tr><td>DOPC</td><td>Avanti</td><td>850375</td><td></td></tr><tr><td>DOPC</td><td>Anatrace</td><td>D518</td><td></td></tr><tr><td>DOPE</td><td>Avanti</td><td>850725</td><td></td></tr><tr><td>DTT</td><td>Fisher</td><td>BP172</td><td></td></tr><tr><td>Dual Thickness MicroLoops</td><td>MiTeGen</td><td></td><td></td></tr><tr><td>EDTA</td><td>BioShop</td><td>EDT003</td><td>Disodium salt, dihydrate</td></tr><tr><td>EGTA</td><td>MilliporeSigma</td><td>324626</td><td></td></tr><tr><td>Emulsifier (EmulsiFex-C3)</td><td>Avestin</td><td></td><td></td></tr><tr><td>Endo H</td><td>New England Biolabs</td><td>P0702</td><td></td></tr><tr><td>Ethanol</td><td>Greenfield</td><td>P016EAAN</td><td>Ethyl Alcohol Anhydrous</td></tr><tr><td>Formaldehyde</td><td>MilliporeSigma</td><td>252549</td><td>ACS Reagent</td></tr><tr><td>Glycerol</td><td>BioShop</td><td>GLY004</td><td></td></tr><tr><td>HEPES</td><td>BioShop</td><td>HEP001</td><td></td></tr><tr><td>HRV-3C protease</td><td></td><td></td><td>Homemade</td></tr><tr><td>Imidazole</td><td>BioShop</td><td>IMD510</td><td>Reagent grade</td></tr><tr><td>Iodoacetamide</td><td>MilliporeSigma</td><td>I1149</td><td>BioUltra</td></tr><tr><td>Isopropanol</td><td>Fisher</td><td>BP2618212</td><td></td></tr><tr><td>Leupeptin</td><td>BioShop</td><td>LEU001</td><td></td></tr><tr><td>MES</td><td>MilliporeSigma</td><td>69892</td><td>BioUltra</td></tr><tr><td>Methanol</td><td>Fisher</td><td>A412P</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Wisent</td><td>800-070-CG</td><td>Hydrated</td></tr><tr><td>microfluidizer (LM 20)</td><td>Microfluidics</td><td></td><td></td></tr><tr><td>NaCl</td><td>BioShop</td><td>SOD002</td><td></td></tr><tr><td>NH&lt;sub&gt;4&lt;/sub&gt;OH</td><td>Fisher</td><td>A669-212</td><td>ACS Reagent</td></tr><tr><td>Ni-NTA superflow</td><td>Qiagen</td><td>30430</td><td>Nickel-charged resins</td></tr><tr><td>PEG 400</td><td>MilliporeSigma</td><td>202398</td><td></td></tr><tr><td>Pepstatin</td><td>BioShop</td><td>PEP605</td><td></td></tr><tr><td>PMSF</td><td>MilliporeSigma</td><td>P7626</td><td></td></tr><tr><td>pSGP18 and pLIC&amp;nbsp;</td><td></td><td></td><td>Homemade (derived from pPICZ, Invitrogen)</td></tr><tr><td>SDS</td><td>BioShop</td><td>SDS003</td><td></td></tr><tr><td>Sodium cholate</td><td>Fisher</td><td>229101</td><td></td></tr><tr><td>Sodium malonate</td><td>MilliporeSigma</td><td>63409</td><td></td></tr><tr><td>Sucrose</td><td>Wisent</td><td>800-081-WG</td><td>Ultra pure</td></tr><tr><td>Superdex 200 30/100 GL</td><td>Cytiva (formerly GE Healthcare Life Sciences)</td><td>28990944</td><td>Prepacked gel-filtration column</td></tr><tr><td>TCEP</td><td></td><td></td><td></td></tr><tr><td>TEM</td><td>FEI, Technai</td><td></td><td></td></tr><tr><td>Tris Base</td><td>Fisher</td><td>BP152</td><td></td></tr><tr><td>&amp;beta;-DDM</td><td>Anatrace</td><td>D310S</td><td>Sol Grade</td></tr><tr><td>&amp;beta;-mercaptoethanol</td><td>MilliporeSigma</td><td></td><td></td></tr><tr><td>&amp;epsilon;-aminocaproic acid</td><td>Fisher</td><td>AAA1471936</td><td></td></tr></tbody></table>

abcg5/g8 crystallization in a lipidic bicelle environment for x-ray crystallography

ATP-binding cassette (ABC) transporters constitute lipid-embedded membrane proteins. Extracting these membrane proteins from the lipid bilayer to an aqueous environment is typically achieved by employing detergents. These detergents disintegrate the lipid bilayer and solubilize the proteins. The intrinsic habitat of membrane proteins within the lipid bilayer poses a challenge in maintaining their stability and uniformity in solution for structural characterization. Bicelles, which comprise a blend of long and short-chain phospholipids and detergents, replicate the natural lipid structure. The utilization of lipid bicelles and detergents serves as a suitable model system for obtaining high-quality diffraction crystals, specifically to determine the high-resolution structure of membrane proteins. Through these synthetic microenvironments, membrane proteins preserve their native conformation and functionality, facilitating the formation of three-dimensional crystals. In this approach, the detergent-solubilized heterodimeric ABCG5/G8 was reintegrated into DMPC/CHAPSO bicelles, supplemented with cholesterol. This setup was employed in the vapor diffusion experimental procedure for protein crystallization.

Recombinant ABC half-transporters, human ABCG5, and ABCG8, are co-expressed in Pichia pastoris yeast. The yeast membrane fraction is then fractionated through centrifugation. As outlined in this protocol, the heterodimeric proteins are extracted using tandem column chromatography. Subsequently, chemically pre-treated proteins are crystallized by incubating them with phospholipid/cholesterol bicelles. Schematic overviews of the purification and crystallization processes are provided in Figure 1.
To assess the monodispersity of the purified proteins, samples containing 0.01-0.05 mg/mL of proteins are stained with 1%-2% uranyl acetate. These samples are then examined using negative-stain TEM (Figure 2A). In order to evaluate protein stability without undergoing freeze-thaw cycles, analytical gel filtration chromatography is employed. This analysis involves monitoring the time-course storage of purified proteins through the use of small, equal-volume aliquots of the proteins (Figure 2B). There might be a slight loss of proteins at the peak fractions after a week of incubation at 4 &#176;C, possibly due to residual soluble protein aggregates. Nonetheless, the overall protein yield remains sufficient for crystal growth. The use of negative stain TEM and analytical gel filtration chromatography is a standard practice to assess the suitability of proteins for crystallization, particularly from different engineered constructs.
For the assessment of protein quality at each step of the column chromatography process, as well as after the pre-crystallization chemical treatment, aliquots of fractions corresponding to two Ni-NTA columns, two CBP columns, one gel filtration, and reductive alkylation are loaded onto a 10% SDS-PAGE gel (Figure 3). Additionally, the same reaction environment utilized for alkylation can be applied for mercury labeling with ethyl mercury (EMTS), although this is beyond the scope of the current study.
The growth of crystals is monitored daily using a tabletop stereo microscope equipped with a polarizer. Crystals that are mature and suitable for data collection generally attain dimensions of 50 &#181;m x 100 &#181;m x 2 &#181;m (Figure 4). During the crystal harvesting process, smaller crystals or clusters are deliberately avoided.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65828/65828fig01.jpg" /> 
Figure 1: Schematic overviews for purification (A) and bicelle crystallization (B) of heterodimeric ABCG5/G8. Constructs of recombinant human ABCG5 (hG5) and ABCG8 (hG8) carry RGS-H6-G-H6 and 3C-CBP tags, respectively (A, top). Tandem affinity column chromatography, followed by gel filtration chromatography to achieve heterodimeric purification (A, bottom). <a href="https://www.jove.com/files/ftp_upload/65828/65828fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65828/65828fig02.jpg" /> 
Figure 2: Evaluation of mono-dispersity (A) and stability (B) of purified proteins. (A) Electron micrograph of negatively stained ABCG5/G8 (G5G8) heterodimers using TEM. Representative particles are highlighted in solid white circles. Scale bar = 100 nm. (B) Alkylated proteins stored at 4 &#176;C analyzed by analytical gel filtration chromatography over the course of a month with a slight loss of proteins after a week. <a href="https://www.jove.com/files/ftp_upload/65828/65828fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65828/65828fig03.jpg" /> 
Figure 3: SDS-PAGE analysis of protein eluates of column chromatography and reductive alkylation. Various volumes (1-10 &#181;L) of protein fractions were loaded onto a 10% Tris/Glycine gel and ran for 45 min at a constant voltage of 200 V. The gel was stained with Coomassie blue, destained, air-dried, and scanned by a tabletop scanner. 1&#176; &#38; 2&#176; Ni: first and second Ni-NTA columns; 1&#176; &#38; 2&#176; CBP: first and second CBP columns; Peak Fractions solid line: pooled fractions for crystallization; Peak Fractions dashed line: shoulder fractions; EMTS: ethyl mercury thiosalicynate; IA: iodoacedamide. <a href="https://www.jove.com/files/ftp_upload/65828/65828fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65828/65828fig04.jpg" /> 
Figure 4: Assessment of protein crystal maturation by light microscopy. Mature crystals of ABCG5/G8 from a crystallization drop were visualized under a tabletop and polarizer-equipped stereo microscope. Scale bar = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/65828/65828fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Bicelle Crystallization Setup for ABC Transporter Membrane Proteins to Advance Drug Development at JoVE.com

Author Spotlight: A Bicelle Crystallization Setup for ABC Transporter Membrane Proteins to Advance Drug Development 

This protocol describes a setup for the crystallization of the sterol transporter ABCG5/G8. ABCG5/G8 is reconstituted into bicelles for hanging-drop crystallization. The protocol does not require specialized materials or substrates, making it accessible and easy to adapt in any laboratory for determining the protein structure through X-ray crystallography.

ABCG5/G8 Crystallization in a Lipidic Bicelle Environment for X-Ray Crystallography

ATP-binding cassette (ABC) transporters constitute lipid-embedded membrane proteins. Extracting these membrane proteins from the lipid bilayer to an ...

abcg5g8-crystallization-lipidic-bicelle-environment-for-x-ray

Research

JoVE Journal

Biochemistry

1.2K Views.  University of Ottawa. This protocol describes a setup for the crystallization of the sterol transporter ABCG5/G8. ABCG5/G8 is reconstituted into bicelles for hanging-drop crystallization. The protocol does not require specialized materials or substrates, making it accessible and easy to adapt in any laboratory for determining the protein structure through X-ray crystallography.

Video: Author Spotlight: A Bicelle Crystallization Setup for ABC Transporter Membrane Proteins to Advance Drug Development 

Preparation and Delivery of Protein Microcrystals in Lipidic Cubic Phase for Serial Femtosecond Crystallography

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural information, typically obtained by crystallography; however MPs are difficult to crystallize. Recent progress in MP structural determination has benefited greatly from the development of lipidic cubic phase (LCP) crystallization methods, which typically yield well-diffracting, but often small crystals that suffer from radiation damage during traditional crystallographic data collection at synchrotron sources. The development of new-generation X-ray free-electron laser (XFEL) sources that produce extremely bright femtosecond pulses has enabled room temperature data collection from microcrystals with no or negligible radiation damage. Our recent efforts in combining LCP technology with serial femtosecond crystallography (LCP-SFX) have resulted in high-resolution structures of several human G protein-coupled receptors, which represent a notoriously difficult target for structure determination. In the LCP-SFX technique, LCP is recruited as a matrix for both growth and delivery of MP microcrystals to the intersection of the injector stream with an XFEL beam for crystallographic data collection. It has been demonstrated that LCP-SFX can substantially improve the diffraction resolution when only sub-10 &#181;m crystals are available, or when the use of smaller crystals at room temperature can overcome various problems associated with larger cryocooled crystals, such as accumulation of defects, high mosaicity and cryocooling artifacts. Future advancements in X-ray sources and detector technologies should make serial crystallography highly attractive and practicable for implementation not only at XFELs, but also at more accessible synchrotron beamlines. Here we present detailed visual protocols for the preparation, characterization and delivery of microcrystals in LCP for serial crystallography experiments. These protocols include methods for conducting crystallization experiments in syringes, detecting and characterizing the crystal samples, optimizing crystal density, loading microcrystal laden LCP into the injector device and delivering the sample to the beam for data collection.

We describe procedures for the preparation and delivery of membrane protein microcrystals in lipidic cubic phase for serial crystallography at X-ray free-electron lasers and synchrotron sources. These protocols can also be applied for incorporation and delivery of soluble protein microcrystals, leading to substantially reduced sample consumption compared to liquid injection.

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural ...

From Constructs to Crystals &#8211; Towards Structure Determination of &#946;-barrel Outer Membrane Proteins

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% percent of known structures. There are two types of membrane proteins, &#945;-helical and &#946;-barrel. While &#945;-helical membrane proteins can be found in nearly all cellular membranes, &#946;-barrel membrane proteins can only be found in the outer membranes of mitochondria, chloroplasts, and Gram-negative bacteria. One common bottleneck in structural studies of membrane proteins in general is getting enough pure sample for analysis. In hopes of assisting those interested in solving the structure of their favorite &#946;-barrel outer membrane protein (OMP), general protocols are presented for the production of target &#946;-barrel OMPs at levels useful for structure determination by either X-ray crystallography and/or NMR spectroscopy. Here, we outline construct design for both native expression and for expression into inclusion bodies, purification using an affinity tag, and crystallization using detergent screening, bicelle, and lipidic cubic phase techniques. These protocols have been tested and found to work for most OMPs from Gram-negative bacteria; however, there are some targets, particularly for mitochondria and chloroplasts that may require other methods for expression and purification. As such, the methods here should be applicable for most projects that involve OMPs from Gram-negative bacteria, yet the expression levels and amount of purified sample will vary depending on the target OMP.

&#946;-barrel outer membrane proteins (OMPs) serve many functions within the outer membranes of Gram-negative bacteria, mitochondria, and chloroplasts. Here, we hope to alleviate a known bottleneck in structural studies by presenting protocols for the production of &#946;-barrel OMPs in sufficient quantities for structure determination by X-ray crystallography or NMR spectroscopy.

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% ...

Chemistry

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Biology

Assessing Two-dimensional Crystallization Trials of Small Membrane Proteins for Structural Biology Studies by Electron Crystallography

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray crystallography to study structure-function questions of membrane proteins, as well as soluble proteins. Screening for two-dimensional (2D) crystals by transmission electron microscopy (EM) is the critical step in finding, optimizing, and selecting samples for high-resolution data collection by cryo-EM. Here we describe the fundamental steps in identifying both large and ordered, as well as small 2D arrays, that can potentially supply critical information for optimization of crystallization conditions.
By working with different magnifications at the EM, data on a range of critical parameters is obtained. Lower magnification supplies valuable data on the morphology and membrane size. At higher magnifications, possible order and 2D crystal dimensions are determined. In this context, it is described how CCD cameras and online-Fourier Transforms are used at higher magnifications to assess proteoliposomes for order and size.
While 2D crystals of membrane proteins are most commonly grown by reconstitution by dialysis, the screening technique is equally applicable for crystals produced with the help of monolayers, native 2D crystals, and ordered arrays of soluble proteins. In addition, the methods described here are applicable to the screening for 2D crystals of even smaller as well as larger membrane proteins, where smaller proteins require the same amount of care in identification as our examples and the lattice of larger proteins might be more easily identifiable at earlier stages of the screening.

Evaluating two-dimensional (2D) crystallization trials for the formation of ordered membrane protein arrays is a highly critical and difficult task in electron crystallography. Here we describe our approach in screening for and identifying 2D crystals of predominantly small membrane proteins in the range of 15 &#x2013; 90kDa.

Electron crystallography has evolved as a method that can be used either alternatively or in combination with three-dimensional crystallization and X-ray ...

Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown into highly ordered two-dimensional (2D) crystals under favorable conditions. The quality of the grown 2D crystals is crucial to the resolution of the final reconstruction via 2D image processing. Over the years, lipid monolayers have been used as a supporting layer to foster the 2D crystallization of peripheral membrane proteins as well as soluble proteins. This method can also be applied to 2D crystallization of integral membrane proteins but requires more extensive empirical investigation to determine detergent and dialysis conditions to promote partitioning to the monolayer. A lipid monolayer forms at the air-water interface such that the polar lipid head groups remain hydrated in the aqueous phase and the non-polar, acyl chains, tails partition into the air, breaking the surface tension and flattening the water surface. The charged nature or distinctive chemical moieties of the head groups provide affinity for proteins in solution, promoting binding for 2D array formation. A newly formed monolayer with the 2D array can be readily transfer into an electron microscope (EM) on a carbon-coated copper grid used to lift and support the crystalline array. In this work, we describe a lipid monolayer methodology for cryogenic electron microscopic (cryo-EM) imaging.

Lipid monolayers have been used as a foundation for forming two-dimensional (2D) protein crystals for structural studies for decades. They are stable at the air-water interface and can serve as a thin supporting material for electron imaging. Here we present the proven steps on preparing lipid monolayers for biological studies.

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown ...

A Package of Established Analytical Tools to Investigate the Solid-State Alteration of Lipid-Based Excipients

Lipid-based excipients (LBEs) are low-toxic, biocompatible, and natural-based, and their application supports the sustainability of pharmaceutical manufacturing. However, the major challenge is their unstable solid-state, affecting the stability of the pharmaceutical product. Critical physical properties of lipids for their processing-such as melt temperature and viscosity, rheology, etc.-are related to their molecular structure and their crystallinity. Additives, as well as thermal and mechanical stress involved in the manufacturing process, affect the solid-state of lipids and thus the performance of pharmaceutical products thereof. Therefore, understanding the alteration in the solid-state is crucial. In this work, the combination of powder x-ray diffraction and differential scanning calorimetry (DSC) is introduced as the gold standard for the characterization of lipids' solid state. X-ray diffraction is the most efficient method to screen polymorphism and crystal growth. The polymorphic arrangement and the lamella length are characterized in the wide- and small-angle regions of x-ray diffraction, respectively. The small-angle x-ray scattering (SAXS) region can be further used to investigate crystal growth. Phase transition and separation can be indicated. DSC is used to screen the thermal behavior of lipids, estimate the miscibility of additives and/or active pharmaceutical ingredients (API) in the lipid matrix, and provide phase diagrams. Four case studies are presented in which LBEs are either used as a coating material or as an encapsulation matrix to provide lipid-coated multiparticulate systems and lipid nanosuspensions, respectively. The lipid solid-state and its potential alteration during storage are investigated and correlated to the alteration in the API release. Qualitative microscopical methods such as polarized light microscopy and scanning electron microscopy are complementary tools to investigate micro-level crystallization. Further analytical methods should be added based on the selected manufacturing process. The structure-function-processability relationship should be understood carefully to design robust and stable lipid-based pharmaceutical products.

This publication shows the application of x-ray diffraction and differential scanning calorimetry as gold standards for investigating the solid-state of lipid-based excipients (LBEs). Understanding the solid-state alteration in LBEs and its effect on the performance of pharmaceutical products thereof is the key factor for manufacturing robust lipid-based dosage forms.

Lipid-based excipients (LBEs) are low-toxic, biocompatible, and natural-based, and their application supports the sustainability of pharmaceutical ...

Protein Crystallization for X-ray Crystallography

Using the three-dimensional structure of biological macromolecules to infer how they function is one of the most important fields of modern biology. The availability of atomic resolution structures provides a deep and unique understanding of protein function, and helps to unravel the inner workings of the living cell. To date, 86% of the Protein Data Bank (rcsb-PDB) entries are macromolecular structures that were determined using X-ray crystallography.
To obtain crystals suitable for crystallographic studies, the macromolecule (e.g. protein, nucleic acid, protein-protein complex or protein-nucleic acid complex) must be purified to homogeneity, or as close as possible to homogeneity. The homogeneity of the preparation is a key factor in obtaining crystals that diffract to high resolution (Bergfors, 1999; McPherson, 1999).
Crystallization requires bringing the macromolecule to supersaturation. The sample should therefore be concentrated to the highest possible concentration without causing aggregation or precipitation of the macromolecule (usually 2-50 mg/ mL). Introducing the sample to precipitating agent can promote the nucleation of protein crystals in the solution, which can result in large three-dimensional crystals growing from the solution. There are two main techniques to obtain crystals: vapor diffusion and batch crystallization. In vapor diffusion, a drop containing a mixture of precipitant and protein solutions is sealed in a chamber with pure precipitant. Water vapor then diffuses out of the drop until the osmolarity of the drop and the precipitant are equal (Figure 1A). The dehydration of the drop causes a slow concentration of both protein and precipitant until equilibrium is achieved, ideally in the crystal nucleation zone of the phase diagram. The batch method relies on bringing the protein directly into the nucleation zone by mixing protein with the appropriate amount of precipitant (Figure 1B). This method is usually performed under a paraffin/mineral oil mixture to prevent the diffusion of water out of the drop.
Here we will demonstrate two kinds of experimental setup for vapor diffusion, hanging drop and sitting drop, in addition to batch crystallization under oil.

The 3-D structure of a molecule provides a unique understanding of how the molecule functions. The principal method for structure determination at near-atomic resolution is X-ray crystallography. Here, we demonstrate the current methods for obtaining three-dimensional crystals of any given macromolecule that are suitable for structure determination by X-ray crystallography.

Using the three-dimensional structure of biological macromolecules to infer how they function is one of the most important fields of modern biology. The ...

Crystallization of Membrane Proteins in Lipidic Mesophases

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.

The protocols describe the essential steps for obtaining diffraction quality crystals of a membrane protein starting from reconstitution of the protein in a lipidic cubic phase (LCP), finding initial conditions with LCP-FRAP pre-crystallization assays, setting up LCP crystallization trials and harvesting crystals.

Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.