We would like to thank Drs. James Bowie and Salem Faham for providing technical expertise and guidance on the bicelle method and Dr. Aviv Paz for useful discussions. We acknowledge Le Du for experimental support. Rachna Ujwal has financial interest in MemX Biosciences LLC, which, however, did not support this work. This work was supported in part by grants from the NIH (RO1 GM078844).

Bicelle based crystallization is comprised of four basic steps (Figure 2): i) preparation of a bicelle forming lipid:amphiphile mixture; ii) incorporation of purified protein into the bicelle medium; iii) crystallization trials (manually or robotically); and iv) visualization, crystal extraction and freezing. These steps are described in detail below
1. Preparation of bicelles
Bicelles can form in a variety of lipid:amphiphile combinations and over a wide range of concentrations. Therefore, an initial composition-based on previous successful conditions- is recommended (Table 1). The most successful mixture is the DMPC:CHAPSO bicelle formulation, which can either be purchased commercially as a premixed ready-to-use formulation (see Table of Reagents below) or prepared in the lab as described. For this exercise we will prepare 1 ml of 35% DMPC:CHAPSO mixture at a 2.8:1 molar ratio.
<ol>
 <li>Weigh out 0.26 g DMPC (Mr 677.9 g/mol), 0.09 g CHAPSO (Mr 630.9 g/mol) and add deionized water to a final volume of 1.0 ml.
 <ul type="disc">
 <li>The bicelle percentage can vary between 10%-40% with a DMPC:CHAPSO molar ratio ranging from 2.6-3.0:1 (Table 1).</li>
 <li>Note: The higher the bicelle concentration the more difficult it is to dissolve the lipid resulting in a higher solution viscosity. However, a concentrated bicelle formulation can be advantageous when the protein concentration is low.</li>
 </ul>
 </li>
 <li>Dissolving the lipid to obtain a homogenous solution requires considerable effort, making this step the most time consuming in the bicelle method. Cycle through the following steps until the DMPC is completely mixed:
 <ol>
 <li>Warm the mixture to ~40&deg;C using a water bath or an incubator and vortex for ~1 minute.
 <ul type="disc">
 <li>Note: As more cycles are carried out, warming the mixture will result in a gel-like consistency making it difficult to vortex.</li>
 </ul>
 </li>
 <li>Cool the mixture on ice and vortex for a few minutes. Cooling helps liquefy the solution making it easier to vortex.
 <ul type="disc">
 <li>Note: As more cycles are performed, the mixture may become cloudy upon cooling.</li>
 </ul>
 </li>
 <li>Repeat the steps listed above (1.2.1 and 1.2.2) until the lipid is completely dissolved.
 <ul type="disc">
 <li>Note: This process may take several hours. Bicelle formation is indicated by the changes in phase behavior of the DMPC:CHAPSO formulation. Upon completion, the mixture will be a clear gel at or above room temperature and a viscous liquid on ice.</li>
 </ul>
 </li>
 </ol>
 </li>
 <li>The bicelle mixture is now ready to use and can be kept at -20&deg;C for long-term storage (up to 5 years). Due to the risk of hydrolysis of the phospholipid head group, it is not advisable to store bicelles at room temperature for extended periods.</li>
</ol>
2. Incorporation of protein into bicelles
Most MP structures obtained from bicelles were crystallized in DMPC:CHAPSO bicelle concentration ranging from 2 to 8% using a protein concentration of 8 to 12 mg/ml (Table 1). If possible, initial screens should use these guidelines and additional concentrations can be screened at the optimization stage. Compared to the LCP method, protein incorporation with bicelles is a simple process (Figure 3), which should be done the same day as crystallization trials.
<ol>
 <li>Thaw the DMPC:CHAPSO bicelle mixture at room temperature until the phase changes to a clear gel.
 <ul type="disc">
 <li>Note: Multiple freeze-thaws will not affect bicelle behavior.</li>
 </ul>
 </li>
 <li>Place the mixture on ice to liquify and briefly vortex to reestablish a homogenous bicelle phase. When placed on ice the mixture may become cloudy.</li>
 <li>From this point on, keep the bicelle mixture and purified protein on ice. This will keep the bicelle in a liquid phase making it amenable to pipetting.</li>
 <li>Add the bicelle mixture to the purified detergent-solubilized protein in a 1:4 (V/V) ratio.
 <ul type="disc">
 <li>For example: 100 &mu;l protein-bicelle mixture is obtained by mixing 80 &mu;l protein with 20 &mu;l bicelle. If the protein concentration is 15 mg/ml and the bicelle concentration is 35%, this will give a bicelle-incorporated protein mixture with a protein concentration of 12 mg/ml and a bicelle concentration of 7%.</li>
 </ul>
 </li>
 <li>Mix by gently pipetting the contents up and down until the solution becomes clear and homogenous.
 <ul type="disc">
 <li>Note: If bubbles appear, a quick spin (30-60 seconds, 13000 rpm, 4&deg;C) with a tabletop centrifuge can help remove them.</li>
 </ul>
 </li>
 <li>Incubate the mixture on ice for at least 30 min to promote complete incorporation of protein into bicelles. The protein-bicelle mixture is now ready for crystallization trials.</li>
</ol>
3. Setting up crystallization trials
Other lipid crystallization techniques like the LCP require specialized equipment due to the medium&prime;s high viscosity; but the unique phase behavior of bicelles permits implementation in virtually any standard crystallization format including robotics (Figure 3). Crystallization trials can be carried out in either hanging or sitting drop formats using standard commercially available screens.
<ol>
 <li>Whether setting up trays manually or using a crystallization robot, maintain the protein-bicelle mixture on ice. This will keep the protein-bicelle mixture cold and ensure that the viscosity of the solution is minimal.</li>
 <li>Manual Crystallization Trials - Using a standard pipette, the protein-bicelle mixture can be mixed with reservoir solution in the same manner as normally performed for soluble or detergent-solubilized membrane proteins.
 <ul type="disc">
 <li>Note: Keep the protein-bicelle mixture on ice during the trials.</li>
 </ul>
 </li>
 <li>Robotic Crystallization Trials - We have tailored these trials for the Mosquito crystallization robot, but in principle (following the same precautions) the technique should be compatible with all crystallization robots. The following tips will ensure that the protein-bicelle mixture remains cool and accurately pipetted by the robot:
 <ol>
 <li>Precool the plate holding the protein-bicelle mixture by placing it on ice. </li>
 <li>Pipette the protein-bicelle mixture into the plate and continue to keep the plate on ice. This plate should be the last item to go on the robot before commencing the run.</li>
 <li>Set-up the reservoir tray and crystallization lids on platform 3 and 5 respectively, of the mosquito robot.</li>
 <li>Place the plate containing protein-bicelle mixture on platform 4 of the mosquito robot. This assures the protein-bicelle mixture is the last to be picked up by the robot and it is immediately released.</li>
 <li>To prevent heating and increased viscosity, do not mix the reservoir with the protein-bicelle mixture.</li>
 <li>When performing multiple screens, immediately return the bicelle-protein plate to ice for cooling as soon as a run is completed.</li>
 </ol>
 </li>
 <li>Drop volume and ratio (protein:reservoir) can be chosen as for conventional crystallization trials. For example, initial crystal trials using the Mosquito robot can be set up using 0.25 &mu;l protein:bicelle mixture plus 0.25 &mu;l reservoir.</li>
 <li>Incubate the crystal trials in a chamber at 20&deg;C. Temperature is a good screening and optimization parameter because the phase behavior of bicelles is temperature dependent.
 <ul type="disc">
 <li>Higher temperatures induce the lamellar phase12 (Figure 1), which has the advantage of pre-organizing the protein in layers. Temperatures below 20&deg;C can be screened, but you should not go below 4&deg;C since this may cause the lipids to precipitate over extended periods of time.</li>
 </ul>
 </li>
 <li>In the same manner as traditional crystallization trials, bicelle trials should be monitored on a regular basis for crystal appearance and growth. We recommend to check trays on the 1st and 3rd day post-setup followed by weekly inspection.</li>
 <li>Crystal optimization can be carried out using methods routinely used for detergent based crystals including grid screening, additive screening, changing temperatures etc. Additionally, bicelle percentage and protein:bicelle ratio can be varied. Furthermore, bicelles can be doped with specific lipids that may be necessary for protein stability or function.</li>
</ol>
4. Visualization, crystal extraction and freezing
Since crystal trials with the protein-bicelle mixture have a viscosity similar to protein-detergent drops, visualization and crystal extraction is routine and is carried out like traditional set-ups.
<ol>
 <li>Visualization: In contrast to LCP media that often requires high-quality illumination under normal and polarized light for crystal detection, visualization is not hindered by bicelles. Colored as well as colorless protein crystal drops can be easily analyzed using standard microscopes and no special equipment is required.</li>
 <li>Bicelles, like other lipidic media, tend to produce a high percentage of false positives. A UV microscope greatly helps in differentiating protein from salt crystals (Figure 4).</li>
 <li>Extraction and Freezing: Crystal extraction and freezing is relatively straightforward and does not require dissolution of the surrounding bicelle media. Additionally, the bicelle phase itself provides some moderate cryo-protection.</li>
</ol>
5. Representative Results:
It generally takes 2-3 days for crystals to appear and approximately a week or more for them to grow to their maximum size. This was the case for bacteriorhodopsin and mouse voltage-dependent anion channel 1 (mVDAC1) crystals4,8. For other membrane proteins it can take several weeks for crystal growth, so it is important to continue monitoring the crystal trials well beyond the first weeks.
As with other lipidic media, bicelles tend to form shapes that may appear to be crystalline. It has also been observed that they lead to a higher percentage of salt and detergent crystals. A UV-microscope that detects tryptophan fluorescence can significantly help eliminate such non-proteinaceous false positives. Figure 4 shows lipid shapes, salt and protein crystals as viewed in visible and UV light to help distinguish the different results that may be observed.
<img alt="Figure 1" src="/files/ftp_upload/3383/3383fig1.jpg" /> 
Figure 1. Bicelle Schematic. Bicelles are composed of a bilayer forming lipid molecule such as DMPC (blue) and an amphiphile such as CHAPSO (green), which protects the hydrophobic edges of the bilayer. As the temperature is increased, disc-like bicelles undergo a phase transformation into a perforated lamellar sheet12.
<img alt="Figure 2" src="/files/ftp_upload/3383/3383fig2.jpg" /> 
Figure 2. Flowchart for the bicelle crystallization method outlining the four basic steps.
<img alt="Figure 3" src="/files/ftp_upload/3383/3383fig3.jpg" /> 
Figure 3. Crystal trials set-up schematic. Purified detergent-solubilized membrane proteins can be directly mixed with bicelles on ice by simply pipetting the contents together. After incubating the protein/bicelle mixture on ice for ~30 minutes, crystallization trials can be set up using any standard format including robotics.
<img alt="Figure 4" src="/files/ftp_upload/3383/3383fig4.jpg" /> 
Figure 4. Visualization of crystal trials. Visible image (top panel) and UV image (bottom panel) of (A) Needle-shaped crystals observed in a salt-only condition. No fluorescence can be detected from the crystals, an indication of false positive. (B) Rod-shaped crystal formed in an MPD condition. The crystal fluoresced weakly but was found to be non-proteinaceous using X-ray diffraction. (C) Crystal observed about four weeks after setting up trials. The strong fluorescence under UV light confirms it is a protein crystal.
<table border="1">
 <tbody>
 <tr>
 <td>No.</td>
 <td>Protein</td>
 <td>Source</td>
 <td>Bicelle Formulation</td>
 <td>Protein Concentration</td>
 <td>Detergent1</td>
 <td>Resolution (&Aring;)</td>
 <td>Reference</td>
 </tr>
 <tr>
 <td>1</td>
 <td>Bacteriorhodopsin2</td>
 <td>Halobacterium salinarum</td>
 <td>8% DMPC:CHAPSO (2.8:1)</td>
 <td>8 mg/ml</td>
 <td>&nbsp;</td>
 <td>2.0</td>
 <td>Faham and Bowie, 2002</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>8% DTPC:CHAPSO (3:1)</td>
 <td>8 mg/ml</td>
 <td>&nbsp;</td>
 <td>1.8</td>
 <td>Faham et al., 2005</td>
 </tr>
 <tr>
 <td>2</td>
 <td>&beta;2-Adrenergic receptor/Fab complex</td>
 <td>Homo sapiens</td>
 <td>8.3% DMPC:CHAPSO (3:1)</td>
 <td>10 mg/ml</td>
 <td>DDM</td>
 <td>3.4/3.7</td>
 <td>Rasmussen et al., 2007</td>
 </tr>
 <tr>
 <td>3</td>
 <td>Voltage-dependent anion channel 1</td>
 <td>Mus musculus</td>
 <td>7% DMPC:CHAPSO (2.8:1)</td>
 <td>12 mg/ml</td>
 <td>LDAO</td>
 <td>2.3</td>
 <td>Ujwal et al., 2008</td>
 </tr>
 <tr>
 <td>4</td>
 <td>Xanthorhodopsin</td>
 <td>Salinibacter ruber</td>
 <td>4.2% DMPC, 5% NM</td>
 <td>4 mg/ml</td>
 <td>DDM</td>
 <td>1.9</td>
 <td>Luecke et al., 2009</td>
 </tr>
 <tr>
 <td>5</td>
 <td>Rhomboid protease</td>
 <td>Escherichia coli</td>
 <td>2% DMPC:CHAPSO (2.6:1)</td>
 <td>9 mg/ml</td>
 <td>Nonyl glucoside</td>
 <td>1.7</td>
 <td>Vinothkumar, 2011</td>
 </tr>
 </tbody>
</table>
1 Detergent used for membrane protein purification 
2 Native lipids from purple membranes may be carried along during purification
Table 1. Summary of crystallization conditions for membrane protein structures solved using bicelles.

<ol>
	<li>Landau, E. M., Rosenbusch, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipidic+cubic+phases:+A+novel+concept+for+the+crystallization+of+membrane+proteins.">Lipidic cubic phases: A novel concept for the crystallization of membrane proteins.</a> Proceedings of the National Academy of Sciences. 93, 14532-14535 (1996).</li><li>Caffrey, M., Lyons, J., Smyth, T., Hart, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+4+Monoacylglycerols:+The+Workhorse+Lipids+for+Crystallizing+Membrane+Proteins+in+Mesophases.">Chapter 4 Monoacylglycerols: The Workhorse Lipids for Crystallizing Membrane Proteins in Mesophases.</a> Current Topics in Membranes. 63, 83-108 (2009).</li><li>Nollert, P., Landau, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Enzymic+release+of+crystals+from+lipidic+cubic+phases.">Enzymic release of crystals from lipidic cubic phases.</a> Biochem. Soc. Trans. 26, 709-713 (1998).</li><li>Cheng, A., Hummel, B., Qiu, H., Caffrey, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple+mechanical+mixer+for+small+viscous+lipid-containing+samples.">A simple mechanical mixer for small viscous lipid-containing samples.</a> Chem. Phys. Lipids. 95, 11-21 (1998).</li><li>Faham, S., Bowie, J. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bicelle+crystallization:+a+new+method+for+crystallizing+membrane+proteins+yields+a+monomeric+bacteriorhodopsin+structure.">Bicelle crystallization: a new method for crystallizing membrane proteins yields a monomeric bacteriorhodopsin structure.</a> J. Mol. Biol. 316, 1-6 (2002).</li><li>Faham, S., Ujwal, R., Abramson, J., Bowie, J. U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+5+Practical+Aspects+of+Membrane+Proteins+Crystallization+in+Bicelles.">Chapter 5 Practical Aspects of Membrane Proteins Crystallization in Bicelles.</a> Current Topics in Membranes. 63, 109-125 (2009).</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, 836-840 (2005).</li><li>Luecke, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Crystallographic+structure+of+xanthorhodopsin,+the+light-driven+proton+pump+with+a+dual+chromophore.">Crystallographic structure of xanthorhodopsin, the light-driven proton pump with a dual chromophore.</a> Proceedings of the National Academy of Sciences. 105, 16561-16565 (2008).</li><li>Ujwal, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+crystal+structure+of+mouse+VDAC1+at+2.3+&#197;+resolution+reveals+mechanistic+insights+into+metabolite+gating.">The crystal structure of mouse VDAC1 at 2.3 &#197; resolution reveals mechanistic insights into metabolite gating.</a> Proceedings of the National Academy of Sciences. 105, 17742-17747 (2008).</li><li>Vinothkumar, K. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+rhomboid+protease+in+a+lipid+environment.">Structure of rhomboid protease in a lipid environment.</a> J. Mol. Biol. 407, 232-247 (2011).</li><li>Rasmussen, S. G. F. <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+[bgr]2+adrenergic+G-protein-coupled+receptor.">Crystal structure of the human [bgr]2 adrenergic G-protein-coupled receptor.</a> Nature. 450, 383-387 (2007).</li><li>Prosser, R. S., Hwang, J. S., Vold, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Magnetically+aligned+phospholipid+bilayers+with+positive+ordering:+a+new+model+membrane+system.">Magnetically aligned phospholipid bilayers with positive ordering: a new model membrane system.</a> Biophys. J. 74, 2405-2418 (1998).</li></ol>

No conflicts of interest declared.

Bicelles are a unique lipidic media that offer a native bilayer-like environment while behaving as if solubilized by detergents. This property gives bicelles a distinct advantage over other lipid-based crystallization methods since there is no learning curve or specialized equipment required for this technique. Once bicelles are available, either commercial or prepared in the lab, they can be directly mixed with purified protein and from this point on crystallization trials proceed almost exactly as with standard deterge...

<table border="1">
	<tbody>
		<tr>
			<td>
				Name of the reagent</td>
			<td>
				Company</td>
			<td>
				Catalogue number</td>
			<td>
				Comments</td>
		</tr>
		<tr>
			<td>DMPC</td>
			<td>Affymetrix</td>
			<td>D514</td>
			<td></td>
		</tr>
		<tr>
			<td>CHAPSO</td>
			<td>Affymetrix</td>
			<td>C317</td>
			<td></td>
		</tr>
		<tr>
			<td>Ready-to-use Bicelles</td>
			<td>MemX Biosciences</td>
			<td>MX201001/MX201002</td>
			<td></td>
		</tr>
		<tr>
			<td>Crystallization Screens</td>
			<td>Qiagen, Hamptop Research, Molecular Dimensions, Emerald Biosystems, Jena Bioscience</td>
			<td></td>
			<td>Standard commercially available screens can be used for initial screening</td>
		</tr>
		<tr>
			<td>Crystallization Set-up</td>
			<td></td>
			<td></td>
			<td>Standard manual and/or robotic set-up available in lab can be used.</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method at JoVE.com

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

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 ...

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23.5K Views.  University of California Los Angeles. 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 tr...

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

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

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 ...

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 ...

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 ...

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.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

RNA Secondary Structure Prediction Using High-throughput SHAPE

Understanding the function of RNA involved in biological processes requires a thorough knowledge of RNA structure. Toward this end, the methodology dubbed "high-throughput selective 2' hydroxyl acylation analyzed by primer extension", or SHAPE, allows prediction of RNA secondary structure with single nucleotide resolution. This approach utilizes chemical probing agents that preferentially acylate single stranded or flexible regions of RNA in aqueous solution. Sites of chemical modification are detected by reverse transcription of the modified RNA, and the products of this reaction are fractionated by automated capillary electrophoresis (CE). Since reverse transcriptase pauses at those RNA nucleotides modified by the SHAPE reagents, the resulting cDNA library indirectly maps those ribonucleotides that are single stranded in the context of the folded RNA. Using ShapeFinder software, the electropherograms produced by automated CE are processed and converted into nucleotide reactivity tables that are themselves converted into pseudo-energy constraints used in the RNAStructure (v5.3) prediction algorithm. The two-dimensional RNA structures obtained by combining SHAPE probing with in silico RNA secondary structure prediction have been found to be far more accurate than structures obtained using either method alone.

High-throughput selective 2' hydroxyl acylation analyzed by primer extension (SHAPE) utilizes a novel chemical probing technology, reverse transcription, capillary electrophoresis and secondary structure prediction software to determine the structures of RNAs from several hundred to several thousand nucleotides at single nucleotide resolution.

Understanding the function of RNA involved in biological processes requires a thorough knowledge of RNA structure. Toward this end, the methodology dubbed ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

Detection of miRNA Targets in High-throughput Using the 3'LIFE Assay

Luminescent Identification of Functional Elements in 3&#8217;UTRs (3&#8217;LIFE) allows the rapid identification of targets of specific miRNAs within an array of hundreds of queried 3&#8217;UTRs. Target identification is based on the dual-luciferase assay, which detects binding at the mRNA level by measuring translational output, giving a functional readout of miRNA targeting. 3&#8217;LIFE uses non-proprietary buffers and reagents, and publically available reporter libraries, making genome-wide screens feasible and cost-effective. 3&#8217;LIFE can be performed either in a standard lab setting or scaled up using liquid handling robots and other high-throughput instrumentation. We illustrate the approach using a dataset of human 3&#8217;UTRs cloned in 96-well plates, and two test miRNAs, let-7c and miR-10b. We demonstrate how to perform DNA preparation, transfection, cell culture and luciferase assays in 96-well format, and provide tools for data analysis. In conclusion 3&#39;LIFE is highly reproducible, rapid, systematic, and identifies high confidence targets.

Detection of miRNA Targets in High-throughput Using the 3&#039;LIFE Assay

Luminescent identification of functional elements in 3&#8217; untranslated regions (3&#8217;UTRs) (3&#8217;LIFE) is a technique to identify functional regulation in 3&#8217;UTRs by miRNAs or other regulatory factors. This protocol utilizes high-throughput methodology such as 96-well transfection and luciferase assays to screen hundreds of putative interactions for functional repression.

Luminescent Identification of Functional Elements in 3&#8217;UTRs (3&#8217;LIFE) allows the rapid identification of targets of specific miRNAs within an ...