Name: crystallization of membrane proteins in lipidic mesophases
Uploaded: 2011-03-28T12:00:00
Duration: 11 min 53 s
Description: Watch this Scientific Journal Video about Crystallization of Membrane Proteins in Lipidic Mesophases at JoVE.com

This work was funded in parts by the NIH grants GM073197 and RR025336.

A typical outline of an in meso crystallization experiment is shown in Fig.11,2. Pre-crystallization LCP-FRAP assays are optional; however, they can significantly accelerate the process of searching for initial crystallization conditions, especially in the case of difficult membrane proteins3.
1. Protein Reconstitution in LCP
<ol>
<li>Purify a membrane protein of interest in a detergent solution and concentrate the protein/detergent complexes to ~10 - 20 mg/mL, taking care not to over-concentrate the detergent1,4.</li>
<li>Transfer ~25 mg of an LCP host lipid (typically monoolein) or a lipid mixture into a 1.5 mL plastic tube and incubate at 40 &deg;C for few minutes until the lipid melts.</li>
<li>Attach a syringe coupler to a 100 &mu;L gas-tight syringe.</li>
<li>Load the syringe with the molten lipid using an adjustable volume pipette. Record the volume of the lipid in the syringe.</li>
<li>Load another 100 &mu;L syringe with the protein solution at a protein solution-to-lipid ratio 2/3 v/v.</li>
<li>Connect both syringes together through the syringe coupler.</li>
<li>Push the syringe plungers alternately to move the lipid and protein through the inner needle of the coupler, back and forth, until the lipid mesophase becomes homogeneous. LCP forms spontaneously upon mechanical mixing, and the protein becomes reconstituted in the lipid bilayer of LCP. Formation of LCP can be verified by its transparent and gel-like consistency and by the absence of birefringency when viewed under a microscope equipped with cross-polarizers, or, if possible, by using small-angle X-ray diffraction1.</li>
</ol>
2. LCP-FRAP Pre-crystallization Assays
LCP-FRAP assays are designed to measure the diffusion properties of membrane proteins reconstituted in LCP at a variety of screening conditions3. The long-range diffusion of membrane proteins in LCP is essential for successful crystallization; however, the microstructure of LCP constrains diffusion of large proteins or oligomeric protein aggregates. A common reason for failure of an in meso crystallization experiment is a fast protein aggregation leading to a loss of diffusion. It has been shown that the aggregation behavior of a protein depends on the particular protein construct, the host lipid and the composition of the screening solution3.
<ol>
<li>Label the protein with a fluorescent dye (Cy3 or similar) at a protein/dye ratio of ~100/1, remove the unreacted dye and concentrate the protein to ~ 1 mg/mL. Label either free amines or free thiols. When labeling free amines, use pH between 7 and 7.5 to predominately label the free N-terminus. Be aware that amino labeling can also label lipids co-purified with the protein2,3.</li>
<li>Reconstitute the labeled protein in LCP as described in section 1).</li>
<li>Set up assay plates as described in section 3) using LCP-FRAP screening solutions instead of crystallization screens2.</li>
<li>Incubate the plates at 20 &deg;C in the dark for at least 12 hours to achieve an equilibrium state.</li>
<li>Place one of the plates on the LCP-FRAP station and focus on the first well using a 10x objective.</li>
<li>Acquire 5 fluorescent images to capture the initial pre-bleached state.</li>
<li>Trigger the laser. The laser power and number of pulses should be adjusted to bleach ~30 - 70% of the labeled protein in the middle of the bleached spot.</li>
<li>Immediately after triggering the laser, start recording a fast post-bleaching sequence of ~200 images at the fastest possible rate.</li>
<li>Follow with recording of a slow post-bleach sequence of ~50 images, selecting the delay between images as 1-20 s, depending on the diffusion rate of the protein.</li>
<li>Integrate the intensity inside the bleach spot in all frames and correct it for bleaching and light intensity fluctuations during the acquisition by dividing the intensity inside the bleached spot by the averaged intensity of a reference spot outside of the laser bleached area.</li>
<li>Normalize the corrected intensity to make the pre-bleached intensity equal to 1 and the initial bleached intensity equal to 0.</li>
<li>Fit the curve of the normalized intensity vs. time, F(t), using the following equation5: 
	F(t) = M x exp(-2T/t) x (I0(2T/t) + I1(2T/t)), (Eq.1) 
where M is the mobile fraction of diffusing molecules, T is the characteristic diffusion time, t is the real time of each recorded frame, I0 and I1 are the 0th and 1st order modified Bessel functions.</li>
<li>Calculate the diffusion coefficient, D, as: 
	D = R2/4T, (Eq.2) 
where R is the radius of the bleached spot.</li>
<li>Move to the next well and repeat steps 2.5) - 2.13).</li>
<li>Compare the mobile fractions and diffusion coefficients obtained for the different screening conditions. Design new crystallization screens based on the components that facilitated protein diffusion and excluding conditions for which protein diffusion was not observed. If the protein did not diffuse in any of the screened conditions, consider broadening the screening space or trying a new protein construct.</li>
</ol>
3. Setting Up LCP Crystallization Trials
<ol>
<li>Reconstitute the protein in LCP as described in section 1).</li>
<li>Transfer the protein-laden LCP into a 10 &mu;L gas-tight syringe attached to a repetitive syringe dispenser.</li>
<li>Attach a short removable needle (gauge 26, 10 mm length) to the 10 &mu;L syringe.</li>
<li>Dispense 200 nL boluses of LCP on the surface of four adjacent wells forming a 2x2 square.</li>
<li>Overlay each of the LCP boluses with 1 &mu;L of corresponding crystallization screen solution.</li>
<li>Cap four loaded wells with an 18 mm square glass coverslip. Apply a gentle pressure on the coverslip to seal the wells.</li>
<li>Repeat steps 3.4)-3.6) with the next set of 4 wells until the whole plate is filled.</li>
<li>Incubate the plate at a constant temperature, periodically checking for crystal formation and growth.</li>
</ol>
4. Harvesting Crystals from LCP
<ol>
<li>Place a plate with protein crystals under a stereo microscope with variable zoom, equipped with a linear rotating polarizer and analyzer.</li>
<li>Focus on the well of interest using a low power zoom so that the whole well is placed within the field of view.</li>
<li>Score the coverslip glass in four strokes making a square inside the well boundaries using a sharp corner of a ceramic capillary cutting stone.</li>
<li>Press around the scored perimeter with strong sharp-point tweezers to propagate the scratches through the thickness of the coverslip glass.</li>
<li>Punch two small holes at opposite corners of the scored square.</li>
<li>Inject few &mu;L of precipitant solution through one of the holes to reduce dehydration during the subsequent steps.</li>
<li>Using an angled sharp needle probe break up the glass along one or two sides to free the cut-out square.</li>
<li>Carefully lift up the glass square watching for the cubic phase bolus. If the bolus is stuck to the coverslip, then flip the glass square over and place on the bottom of the well.</li>
<li>Add an extra few &mu;L of precipitant solution, supplemented with a cryo-protectant, if necessary, on top of the exposed cubic phase bolus in the well.</li>
<li>Increase magnification of the microscope and focus on a crystal.</li>
<li>Adjust the angle between the polarizer and the analyzer to increase the contrast between the birefringent crystal and the background, while keeping enough light to see the harvesting loop.</li>
<li>Select a MiTeGen MicroMount with a diameter matching the crystal size and then harvest the crystal directly from the LCP by scooping it into the MicroMount.</li>
<li>Flash freeze the MicroMount with the harvested crystal in liquid nitrogen, and ship it to a synchrotron source beamline for X-ray data collection6.</li>
</ol>
5. Representative Results:
An engineered human beta 2 adrenergic G protein-coupled receptor (&beta;2AR-T4L) was expressed in baculovirus infected sf9 insect cells and purified in dodecylmaltoside (DDM)/ cholesteryl hemisuccinate (CHS) detergent solution bound to a partial inverse agonist carazolol7. The protein was labeled with Cy3 NHS ester and used in LCP-FRAP pre-crystallization assays (Figure 2). Coarse grid screens based on several conditions selected from the results of LCP-FRAP assays produced initial crystal-like hits (Figure 3). Further optimization of precipitant conditions yielded diffraction quality crystals (Figure 4).
<img src="/files/ftp_upload/2501/2501fig1.jpg" alt="Figure 1" /> 
Figure 1. Flow-chart of a typical LCP crystallization experiment. Steps in the gray boxes are not described in the current protocols.
<img src="/files/ftp_upload/2501/2501fig2.jpg" alt="Figure 2" /> 
Figure 2. LCP-FRAP assay with &beta;2AR-T4L/carazolol in monoolein based LCP. A) Results of an LCP-FRAP assay performed in an automatic high-throughput mode, in which each sample of a 96-well plate is bleached sequentially and fluorescence recovery is measured after a 30 min incubation. The obtained fluorescence recoveries, which represent the mobile fraction in each sample, are plotted for all 96 samples. The screening solutions contain 0.1 M Tris pH 8, 30 % v/v PEG 400 combined with 48 different salts at two different concentrations. B) Fluorescence recovery profiles for several representative conditions. Solid line curves represent fits by Eq. 1.The mobile fractions and the diffusion coefficients are determined using Eqs. 1 and 2. Fast recovery of less than 10% in the sample containing Na chloride is due to fluorescently labeled lipids co-purified with the protein.
<img src="/files/ftp_upload/2501/2501fig3.jpg" alt="Figure 3" /> 
Figure 3. Initial crystal hits of &beta;2AR-T4L/carazolol obtained by a coarse grid screening around most promising conditions identified by LCP-FRAP, containing Na sulfate (panel A) and Na Formate (panel B). The protein is labeled with Cy3 NHS ester and the fluorescent images are taken using excitation at 543 nm and emission at 605 nm. 
<img src="/files/ftp_upload/2501/2501fig4.jpg" alt="Figure 4" /> 
Figure 4. Optimized crystals of &beta;2AR-T4L/carazolol. The images of crystals grown in the presence of Na sulfate (panels A and B) and K Formate (panels C and D) are taken in the brightfield mode (panels A and C) and using cross-polarizers (panels B and D).

<ol>
	<li>Caffrey, M., Cherezov, V. <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+mesophases.">Crystallizing membrane proteins using lipidic mesophases.</a> Nat. Protoc. 4, 706-731 (2009).</li><li>Cherezov, V., Abola, E., Stevens, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+progress+in+the+structure+determination+of+GPCRs,+a+membrane+protein+family+with+high+potential+as+pharmaceutical+targets.">Recent progress in the structure determination of GPCRs, a membrane protein family with high potential as pharmaceutical targets.</a> Methods Mol. Biol. 654, 141-170 (2010).</li><li>Cherezov, V., Liu, J., Hanson, M. A., Griffith, M. T., Stevens, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=LCP-FRAP+assay+for+pre-screening+membrane+proteins+for+in+meso+crystallization.">LCP-FRAP assay for pre-screening membrane proteins for in meso crystallization.</a> J. Cryst. Growth Design. 8, 4307-4315 (2008).</li><li>Misquitta, Y., 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=Detergents+destabilize+the+cubic+phase+of+monoolein:+implications+for+membrane+protein+crystallization.">Detergents destabilize the cubic phase of monoolein: implications for membrane protein crystallization.</a> Biophys. J. 79, 394-405 (2003).</li><li>Soumpasis, D. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Theoretical+analysis+of+fluorescence+photobleaching+recovery+experiments.">Theoretical analysis of fluorescence photobleaching recovery experiments.</a> Biophys. J. 41, 95-97 (1983).</li><li>Cherezov, V., Hanson, M. A., Griffith, M. T., Hilgart, M. C., Sanishvili, R., Nagarajan, V., Stepanov, S., Fischetti, R. F., Kuhn, P., Stevens, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rastering+strategy+for+screening+and+centering+of+microcrystal+samples+of+human+membrane+proteins+with+a+sub-10+micrometer+size+X-ray+synchrotron+beam.">Rastering strategy for screening and centering of microcrystal samples of human membrane proteins with a sub-10 micrometer size X-ray synchrotron beam.</a> J. R. Soc. Interface. 6, S587-S597 (2009).</li><li>Cherezov, V., Rosenbaum, D. M., Hanson, M. A., Rasmussen, S. G., Thian, F. S., Kobilka, T. S., Choi, H. J., Kuhn, P., Weis, W. I., Kobilka, B. K., Stevens, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+crystal+structure+of+an+engineered+human+beta2-adrenergic+G+protein-coupled+receptor.">High-resolution crystal structure of an engineered human beta2-adrenergic G protein-coupled receptor.</a> Science. 318, 1258-1265 (2007).</li><li>Cherezov, V., Peddi, A., Muthusubramaniam, L., Zheng, Y. F., 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+robotic+system+for+crystallizing+membrane+and+soluble+proteins+in+lipidic+mesophases.">A robotic system for crystallizing membrane and soluble proteins in lipidic mesophases.</a> Acta Crystallogr. D. 60, 1795-1807 (2004).</li><li>Kissick, D. J., Gualtieri, E. J., Simpson, G. J., Cherezov, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonlinear+optical+imaging+of+integral+membrane+protein+crystals+in+lipidic+mesophases.">Nonlinear optical imaging of integral membrane protein crystals in lipidic mesophases.</a> Anal. Chem. 82, 491-497 (2010).</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>Cherezov, V., 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+and+inexpensive+nanoliter-volume+dispenser+for+highly+viscous+materials+used+in+membrane+protein+crystallization.">A simple and inexpensive nanoliter-volume dispenser for highly viscous materials used in membrane protein crystallization.</a> J. Appl. Cryst. 38, 398-400 (2005).</li><li>Cherezov, V., 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=Nano-volume+plates+with+excellent+optical+properties+for+fast,+inexpensive+crystallization+screening+of+membrane+proteins.">Nano-volume plates with excellent optical properties for fast, inexpensive crystallization screening of membrane proteins.</a> J. Appl. Cryst. 36, 1372-1377 (2003).</li><li>Cherezov, V., Fersi, 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=Crystallization+screens:+compatibility+with+the+lipidic+cubic+phase+for+in+meso+crystallization+of+membrane+proteins.">Crystallization screens: compatibility with the lipidic cubic phase for in meso crystallization of membrane proteins.</a> Biophys J. 81, 225-242 (2001).</li></ol>

No conflicts of interest declared.

The protocols provide a basic visual guidance for the main steps involved in performing in meso crystallization experiments. More in-depth details related to these protocols, emphasizing possible pitfalls, shortcomings or alternative routes are available elsewhere1,2. Optional LCP-FRAP assays can help at the earlier stages to select the most promising protein construct, LCP host lipid and lipid additives, as well as limit the range of possible precipitants and buffer conditions3. Once an initial crystallization hit is found, it should be optimized to obtain better quality crystals. Optimization of in meso crystallization conditions is essentially similar to optimizing conditions for soluble proteins with the addition of extra parameters associated with the composition of LCP1. Membrane protein crystals grown in lipidic mesophase are typically smaller in size than crystals obtained in detergent solution, but more ordered, thus, benefitting strongly from using microfocus beamlines available at modern synchrotron sources6.
Many of the procedures related to in meso crystallization, including setting up crystallization or assay plates, conducting LCP-FRAP assays and detecting crystals, have been semi- or fully automated1,2,8,9, allowing screening of a large range of conditions while consuming small amounts of protein and lipid. On the other hand, protein reconstitutions in LCP and crystal harvesting remain manual and more tedious operations and, thus, have a need for improvement.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>100 &mu;L gas-tight syringe</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>7656-01</td>
<td>2 syringes are required</td>
</tr>
<tr>
<td>10 &mu;L gas-tight syringe</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>7653-01</td>
<td>&nbsp;</td>
<tr>
<td>Syringe coupler</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>7770-02 30902</td>
<td>The coupler can be made using available parts as described in refs. 10</td>
</tr>
<tr>
<td>Repetitive syringe dispenser</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>83700</td>
<td>The repetitive syringe dispenser can be modified to reduce dispensing volume by ~3 times11</td>
</tr>
<tr>
<td>Short (0.375&#x201D;) flat-tipped removable needle (point style 3, gauge 26)</td>
<td>&nbsp;</td>
<td>Hamilton</td>
<td>7804-03</td>
<td>&nbsp;</td>
<tr>
<td>96-well glass sandwich plate</td>
<td>&nbsp;</td>
<td>Marienfeld</td>
<td>08 900 03</td>
<td>For manual operations it is more convenient to assemble the glass sandwich plate using a standard microscope glass slides and a double sticky tape with punched holes1,2,12.</td>
</tr>
<tr>
<td>Glass cover slip</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>63787-01</td>
<td>&nbsp;</td>
<tr>
<td>monoolein</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M7765</td>
<td>&nbsp;</td>
<tr>
<td>Crystallization screens</td>
<td>&nbsp;</td>
<td>Hampton Research, Molecular Dimensions, Emerald Biosystems, Jena Bioscience</td>
<td>&nbsp;</td>
<td>Most of available commercial screens can be used for initial screening. Conditions that consistently disrupt LCP can be diluted 2x for better compatibility13.</td>
</tr>
<tr>
<td>Capillary cutting stone</td>
<td>&nbsp;</td>
<td>Hampton Research</td>
<td>HR4-334</td>
<td>&nbsp;</td>
<tr>
<td>Fine point tweezers</td>
<td>&nbsp;</td>
<td>Ted Pella</td>
<td>510</td>
<td>&nbsp;</td>
<tr>
<td>Angled sharp probe</td>
<td>&nbsp;</td>
<td>Ted Pella</td>
<td>13650</td>
<td>&nbsp;</td>
<tr>
<td>MicroMounts</td>
<td>&nbsp;</td>
<td>MiTeGen</td>
<td>M1-Lxx-xx</td>
<td>Select MicroMount diameter to match the crystal size</td>
</tr>		</tbody>
		</table>
		</div>

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.

Watch this Scientific Journal Video about Crystallization of Membrane Proteins in Lipidic Mesophases at JoVE.com

Crystallization of Membrane Proteins in Lipidic Mesophases

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 ...

crystallization-of-membrane-proteins-in-lipidic-mesophases

Research

JoVE Journal

Biology

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 ...

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2, ion pumps3, and transporters4. Recent developments have combined site-specific fluorophore labeling alongside TEVC to 
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8. 
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4. Finally, recent developments 
have also enabled the manipulation of select internal ions following co-expression with a second protein9.
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (&lt;5%)3 upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of 
the membrane protein in order to correlate the conformational dynamics to the function of the protein10. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.

Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling

We will describe a method which measures the kinetics of ion transport of membrane proteins alongside site-specific analysis of conformational changes using fluorescence on single cells. This technique is adaptable to ion channels, transporters and ion pumps and can be utilized to determine distance constraints between protein subunits.

Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane ...

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.

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 ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and ...

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and the inherent instability of many membrane proteins once solubilized in detergents. A protocol is described that combines ligation independent cloning of membrane proteins as GFP fusions with expression in Escherichia coli detected by GFP fluorescence. This enables the construction and expression screening of multiple membrane protein/variants to identify candidates suitable for further investment of time and effort. The GFP reporter is used in a primary screen of expression by visualizing GFP fluorescence following SDS polyacrylamide gel electrophoresis (SDS-PAGE). Membrane proteins that show both a high expression level with minimum degradation as indicated by the absence of free GFP, are selected for a secondary screen. These constructs are scaled and a total membrane fraction prepared and solubilized in four different detergents. Following ultracentrifugation to remove detergent-insoluble material, lysates are analyzed by fluorescence detection size exclusion chromatography (FSEC). Monitoring the size exclusion profile by GFP fluorescence provides information about the mono-dispersity and integrity of the membrane proteins in different detergents. Protein: detergent combinations that elute with a symmetrical peak with little or no free GFP and minimum aggregation are candidates for subsequent purification. Using the above methodology, the heterologous expression in E. coli of SED (shape, elongation, division, and sporulation) proteins from 47 different species of bacteria was analyzed. These proteins typically have ten transmembrane domains and are essential for cell division. The results show that the production of the SEDs orthologues in E. coli was highly variable with respect to the expression levels and integrity of the GFP fusion proteins. The experiment identified a subset for further investigation.

Green Fluorescent Protein-based Expression Screening of Membrane Proteins in Escherichia coli

A streamlined approach to screening for the expression of recombinant membrane proteins in Escherichia coli based on fusion to green fluorescent protein is presented.

The production of recombinant membrane proteins for structural and functional studies remains technically challenging due to low levels of expression and ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...