There are many who contributed to this work and most are from the Caffrey Membrane Structural and Functional Biology Group, both past and present members. To all we extend our warmest thanks and appreciation. This work was supported in part by grants from Science Foundation Ireland (07/IN.1/B1836), the National Institutes of Health (GM75915), and the University of Limerick.

An important focus in the area of structural and functional biology is the biological membrane (Figure 1). The membrane, which surrounds the cell and sub-cellular organelles when present, is a molecularly thin structure just two lipid molecules across and is studded with proteins. Structure and function as applied to both lipid and proteins are of interest. However, the focus of this article is restricted to membrane proteins.
A better understanding of how membrane proteins function at a molecular level is sought for two reasons. Firstly, there is the intellectual satisfaction in knowing how they work. Secondly, by knowing how a protein works, there is always the prospect of being able to fix it should it malfunction or of improving or even modifying it for particular applications. Drug design is an obvious outcome of this type of work. One approach to figuring how a membrane protein works at a molecular level is to determine its structure. This involves establishing the location in 3-dimensional space of all atoms, or at least all non-hydrogen atoms, that make up the protein. The method we use for this purpose is macromolecular X-ray crystallography (MX). Figure 2 shows an example of a membrane protein whose structure was determined using MX. A diffraction quality crystal of the protein is required to do MX.
Clearly, there are many steps involved in structure determination using macromolecular crystallography. This is illustrated in Figure 3. Typically, these include identifying a membrane protein target, and then producing, purifying and crystallizing it. Diffraction measurements are performed on the crystal using a home or a synchrotron X-ray source. The diffraction data are processed yielding an electron density map that is then fitted with a molecular model. The model, when refined, can be used to explore the mechanism of action of the protein and for structure-based drug design.
The focus of this article is to show how we produce diffraction quality crystals of membrane proteins using lipidic mesophases, by the so-called in meso method. A recent review of the method and its scope is available in Reference 1 (Caffrey, 2009). The step-by-step protocol we will follow here is described in Reference 2 (Caffrey and Cherezov, 2009).
A flowchart summarizing the steps involved in and time required for setting up an in meso membrane protein crystallization trial is shown in Figure 4. This article covers those steps enclosed by dashed red lines.
Part 1: Preparing the Crystallization Plates
<ol>
 <li>Position a silanized microscope slide on the work bench.</li>
 <li>Remove the protective paper cover from one surface of a strip of perforated double stick spacer tape (commercially available, see Table of Specific Reagents and Equipment below).</li>
 <li>Place the tape, sticky side down, in contact with the surface of the slide.</li>
 <li>Pressure-seal the tape to the slide using a brayer or roller. Aluminum foil can be placed between the tape and the roller to protect the base of the wells. </li>
 <li>Remove the second paper cover from the tape to expose its upper sticky surface.</li>
 <li>Place individual silanized coverslips in close proximity to the plate for fast and efficient sealing of wells as soon as loading is complete.</li>
</ol>
Part 2: Preparing the Lipid Syringe
<ol>
 <li>Place the lipid (often monoolein) in a temperature-controlled block at 45 &deg;C for 3 minutes to render it molten.</li>
 <li>While the lipid is melting prepare two 100-&mu;L gas-tight Hamilton syringes for use in the lipid-protein mixing step. Remove the Teflon ferrule from the syringe that will contain the protein solution. Place the syringe that will hold the lipid next to it (leaving its ferrule in place).</li>
 <li>Connect the coupler (see Table of Specific Reagents and Equipment below and Reference 3 (Cheng et al., 1998)) to the lipid syringe. Finger-tighten the coupler to the syringe but do not over-tighten. Remove the plunger from the barrel of the lipid syringe.</li>
 <li>Note that the lipid must be molten before proceeding. Set the pipette to 30 &mu;L and slowly take up approximately 30 &mu;L of molten lipid. Slowly deliver as much of the molten lipid as you can into the open end of the syringe taking care not to introduce air gaps.</li>
 <li>Place the plunger into the barrel in contact with the molten lipid and slowly advance the plunger up the barrel with the syringe held vertically as demonstrated in the video. Air bubbles that may be trapped inadvertently should rise in the process and be released.</li>
 <li>Carefully determine the volume of lipid in the syringe by reading the markings on the syringe barrel.</li>
 <li>Slide the ferrule onto the needle extending from the coupler. Use the plunger to force the molten lipid up the barrel and slowly and gently into the needle at the core of the coupler.</li>
</ol>
Part 3: Preparing the Protein Syringe
<ol>
 <li>The protein solution should be spun at 14,000 g for 5 to 10 minutes at 4 &deg;C to remove large aggregates before setting up crystallization trials. Remove the protein solution from the ice and allow it to equilibrate at room temperature. </li>
 <li>Calculate the volume of protein solution to use to form the cubic phase at or close to full hydration. For monoolein at 20 &deg;C, full hydration with water occurs at close to 40 % water (by mass) as in the phase diagram5 (Figure 5).</li>
 <li>With a 25 or 50 &mu;L Hamilton syringe take up the required volume of protein solution. Transfer the solution onto the Teflon tip of the plunger in the barrel of the protein syringe taking care to avoid air bubbles.</li>
 <li>Carefully withdraw the needle and measure the volume of protein solution in the syringe. It should match the value delivered in the previous step.</li>
 <li>Slowly inch the protein solution up the barrel to end up flush with its open end.</li>
 <li>Screw the protein syringe into the open end of the coupler attached to the lipid syringe. It is critical that the device not be excessively tightened. Over-tightening the assembled mixing device at this stage will damage the ferrules and cause leaking. Under-tightening will also lead to leaking.</li>
</ol>
Part 4: Mixing Protein Solution and Lipid: Making the Mesophase
<ol>
 <li>To effect mixing, advance the plunger on the protein side of the assembled mixing unit to its limit, with the thumb or index finger driving the protein solution out of the protein syringe, through the coupler and into the lipid syringe. </li>
 <li>The plunger on the lipid side is now used to drive the contents of the lipid syringe back through the coupler and into the protein syringe.</li>
 <li>The process is repeated many times; occasionally, a hundred passages or more are required to produce a homogenous mesophase. At the start of mixing, movement of material back and forth through the coupler can be uneven and, at times, extra force is needed to effect mixing. Initial mixing is usually accompanied by the development of a non-uniform cloudiness in the sample. As homogenization progresses, the texture becomes more uniform and characteristic of the viscous cubic phase, as does the visual appearance of the emerging cubic mesophase. </li>
 <li>If conditions are appropriate and the cubic phase forms, the dispersion should appear optically transparent in the syringe barrel. Thus, the markings on the syringe barrel should be clearly legible through the mesophase in the barrel.</li>
 <li>Very slight cooling of the sample during mixing by placing the syringe mixer for a short time on ice can accelerate homogenization and the achievement of transparency. However, it is important not to overcool the sample.</li>
 <li>Care should be taken to avoid extremely vigorous mixing, as this can cause sample temperature to rise due to frictional heating3.</li>
</ol>
Part 5: Loading the Dispenser
<ol>
 <li>Remove the needle and the plunger from a 10-&mu;L Hamilton syringe. Leave the Teflon ferrule in place.</li>
 <li>Remove the retaining nut from the dispenser with the aid of a small coin.</li>
 <li>With the ratchet arm fully withdrawn, insert the syringe - without its plunger and needle - through the holding ring of the dispenser.</li>
 <li>Replace the retaining nut, gasket side facing the syringe, and screw it in tightly on the open end of the syringe. 
 Care should be taken to ensure the syringe is properly centered in the holding ring and that the barrel is aligned parallel to the ratchet arm. Both can be judged by eye. These two requirements are critical to avoid leakage.</li>
 <li>Pass the plunger through the gripping ring with the gripper nut unscrewed a few turns and guide the plunger into the open end of the syringe. Depress the ratchet arm fully. Move the plunger into the barrel and check that it travels freely and true in the barrel. It may help to loosen the screw on the guide bar to facilitate the plunger moving freely. Make sure that it does. Depress the plunger until its Teflon tip aligns with the zero &mu;L graduation mark on the barrel. If the screw on the guide bar was loosened, retighten it and recheck that the plunger moves freely.</li>
 <li>Move the plunger on one side of the assembled mixing unit to the zero &mu;L graduation mark to transfer the mesophase to the other syringe and the coupler.</li>
 <li>Disconnect the empty syringe (with its ferrule in place) from the mixing unit and immediately connect the loaded syringe - with the coupler attached - to the threaded termination of the 10-&mu;L dispensing syringe. The degree of tightness with which coupling is done is critical, as already noted.</li>
 <li>Load the dispensing syringe by depressing the plunger of the 100-&mu;L syringe so that the mesophase transfers through the coupler.</li>
 <li>Secure a short, flat-tipped needle to the steel termination of the dispensing syringe and carefully tighten it in place.</li>
 <li>Clamp the plunger to the ratchet arm by tightening the nut in the gripping ring. Care should be taken not to over-tighten the nut, as this can score and deform the plunger rendering it unusable. It is important that the travel range provided to the ratchet arm in the clamped condition be limited to no more than an inch, as demonstrated in the video. Beyond this, the plunger will have a tendency to buckle and delivery can fail.</li>
 <li>Depress the ratchet drum several times to advance the plunger in the barrel, thereby filling the void volume of the needle and loading the needle. This step should be repeated (up to ten times) until a continuous string of the mesophase emerges from the tip of the needle.</li>
 <li>Crystallization trials should commence immediately, as some proteins are unstable in the cubic phase without added precipitant.</li>
</ol>
Part 6: Setting up Crystallization Plates
<ol>
 <li>Place a multi-well crystallization plate and coverslip on a surface raised a few inches above the work bench for ease of loading. Optimal contrast and enhanced visibility are achieved when the surface is slightly dark.</li>
 <li>Homogenize the precipitant solutions and uncap the vials. Set the precipitant dispensing pipet to 1 &mu;L.</li>
 <li>With the dispensing syringe held vertically in one hand, use the free hand to position the needle tip in the center and directly above the base of well number 1. Press the button on the repeating dispenser to expel a bolus of mesophase onto the glass surface. The bolus volume is 200 nL when the standard repeating dispenser is used with a 10-&mu;L syringe. The tip of the needle should be no more than a few hundred micrometers above the base of the well to ensure proper delivery.</li>
 <li>After 4 adjacent wells are loaded with mesophase, place 1 &mu;L of precipitant solution on top of each mesophase bolus using a 2-&mu;L pipet and standard disposable tips.</li>
 <li>As quickly as possible, place a coverslip squarely over the filled wells to cover them uniformly. To effect a water-tight seal, use a spatula to apply pressure to the coverslip where it makes contact with the exposed sticky surface of the spacer tape.</li>
 <li>The mesophase and precipitant dispensing process can be repeated until all wells on the plate are loaded and sealed.</li>
 <li>Label the plate clearly for tracking purposes. Light-sensitive proteins are usually handled by wrapping the plates in aluminum foil before placing them in the incubation chamber.</li>
 <li>Place the plates in a temperature-controlled chamber, usually at 20 &deg;C.</li>
 <li>On a regular schedule, inspect the wells for crystal growth using a Polarized Light Microscope with a 10 or 20-fold objective. The schedule used in the author's lab is as follows: day 0, 1, 2, 3, 5, 7, 14, 21, 30 and 60 post-setup. Carefully inspect the mesophase bolus adjusting the depth of focus within the 0.14-mm-thick sample. Examination should be done both in normal light and between crossed-polarizers.</li>
</ol>
Part 7: Representative Results
The appearance of the resulting crystals will vary with the inherent color of the membrane protein, the polarization of the light used for inspection (or lack thereof) and the method and quality of illumination. Figure 6 shows several possible crystal appearances. Naturally colored membrane proteins growing in meso when viewed with normal light can look like those shown in Figure 6 (Panels b and d). Colorless membrane protein crystals growing in the cubic phase when viewed with normal light can look like those shown in Figure 6 (Panel e). Finally, colorless membrane protein crystals growing in meso when viewed with polarized light can look like Figure 6 (Panels a and c).
The next steps in the overall process of structure determination are to harvest and cryo-cool the crystals and to record and analyze X-ray diffraction from them. These topics are to be covered in separate JoVE articles.
<img src="/files/ftp_upload/1712/1712fig1.jpg" alt="Figure 1" /> 
 Figure 1. Schematic representation of a biological membrane showing the lipid bilayer in and on which are situated a variety of proteins.
<img src="/files/ftp_upload/1712/1712fig2.jpg" alt="Figure 2" /> 
 Figure 2. The structure of the vitamin B12 transporting protein, BtuB, solved using MX and crystals grown by the in meso method6 illustrated in this JoVE article.
<img src="/files/ftp_upload/1712/1712fig3.jpg" alt="Figure 3" /> 
 Figure 3. The structure-function cycle illustrates many of the steps involved in obtaining and utilizing complete structural information about a protein.
<img src="/files/ftp_upload/1712/1712fig4.jpg" alt="Figure 4" /> 
 Figure 4. The flowchart summarizes the steps involved in the production of membrane protein crystals by the in meso method. Only those steps surrounded by the dashed red line are covered in this JoVE article. From Reference 2.
<img src="/files/ftp_upload/1712/1712fig5.jpg" alt="Figure 5" /> 
 Figure 5. A simplified temperature-composition phase diagram for the lipid (monoolein)/water system. Crystallization trials are set up at 20 &deg;C where the lipid saturates with water at 40 % hydration. The detailed phase diagram is available in Reference 5.
<img src="/files/ftp_upload/1712/1712fig6.jpg" alt="Figure 6" /> 
 Figure 6. Crystals of membrane proteins growing in the lipidic mesophase. 
 (a) vitamin B12 transporting protein, BtuB6, (b) light-harvesting complex II7, (c) the adhesin/invasin OpcA8, (d) bacteriorhodopsin9, (e) a carbohydrate transporter from Pseudomonas. Images recorded with normal light (b,d,e) and between crossed polarizers (a,c).

<ol>
	<li>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=Crystallizing+membrane+proteins+for+structure+determination.+Use+of+lipidic+mesophases.">Crystallizing membrane proteins for structure determination. Use of lipidic mesophases.</a> Annu. Rev. Biophys. 38, 29-51 (2009).</li><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=Crystallising+membrane+proteins+in+lipidic+mesophases.">Crystallising membrane proteins in lipidic mesophases.</a> Nat. Protoc. 4, 706-731 (2009).</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=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>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=The+phase+diagram+of+the+monoolein/water+system:+metastability+and+equilibrium+aspects.">The phase diagram of the monoolein/water system: metastability and equilibrium aspects.</a> Biomaterials. 21, 223-234 (2000).</li><li>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=In+meso+structure+of+the+cobalamin+transporter,+BtuB,+at+1.95+A+resolution.">In meso structure of the cobalamin transporter, BtuB, at 1.95 A resolution.</a> J. Mol. Biol. 364, 716-734 (2006).</li><li>Cherezov, V., Clogston, J., Papiz, M. Z., 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=Room+to+move:+crystallizing+membrane+proteins+in+swollen+lipidic+mesophases.">Room to move: crystallizing membrane proteins in swollen lipidic mesophases.</a> J. Mol. Biol. 357, 1605-1618 (2006).</li><li>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=In+meso+crystal+structure+and+docking+simulations+suggest+an+alternative+proteoglycan+binding+site+in+the+OpcA+outer+membrane+adhesin.">In meso crystal structure and docking simulations suggest an alternative proteoglycan binding site in the OpcA outer membrane adhesin.</a> Proteins. 71, 24-34 (2008).</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 Biol. Crystallogr. 60, 1795-1807 (2004).</li></ol>

The cubic phase is a delicate and dynamic environment that can change drastically with the alteration of a number of variables. It is not possible to give a description of the setup of in meso crystallization trials in the manual mode that describes all potential pitfalls. However, many difficulties can be avoided by practicing the technique before applying it to expensive protein solutions and by using moderation in pressure applied to syringes during mixing. Done correctly, the in meso method can yield crystals of a wide variety of proteins, the number of which is constantly increasing. The description given here of the setup of in meso crystallization trials is focused on the manual mode. The process can be, and often is modified to facilitate automated setup of the crystallization plates in those cases that require large-scale screening of crystallization conditions.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<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>Protein solution</td>
<td>Reagent</td>
<td>Various</td>
<td>Various</td>
<td>&nbsp;</td>
<tr>
<td>Lipid (monoolein)</td>
<td>Reagent</td>
<td>Sigma/NuCheck</td>
<td>Various</td>
<td>&nbsp;</td>
<tr>
<td>Precipitant solutions</td>
<td>Reagent</td>
<td>Various</td>
<td>Various</td>
<td>&nbsp;</td>
<tr>
<td>Purified water</td>
<td>Reagent</td>
<td>Millipore</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Pipetting devices</td>
<td>Tool</td>
<td>Pipetman</td>
<td>Various</td>
<td>&nbsp;</td>
<tr>
<td>Disposable pipet tips</td>
<td>Disposable</td>
<td>Pipetman</td>
<td>Various</td>
<td>&nbsp;</td>
<tr>
<td>Gas-tight syringes</td>
<td>Tool</td>
<td>Hamilton</td>
<td>80265 (25-&mu;l)</td>
<td>&nbsp;</td>
<tr>
<td>Syringe tips</td>
<td>Tool</td>
<td>Hamilton</td>
<td>7770-020 (gauge 22)</td>
<td>&nbsp;</td>
<tr>
<td>Narrow bore coupler*</td>
<td>Tool</td>
<td>Hamilton/Emerald</td>
<td>various/EBS-LCP-2</td>
<td>&nbsp;</td>
<tr>
<td>Repeating dispenser</td>
<td>Tool</td>
<td>Hamilton</td>
<td>83700</td>
<td>&nbsp;</td>
<tr>
<td>Silanized glass microscope slides</td>
<td>Disposable</td>
<td>GoldSeal</td>
<td>3010</td>
<td>&nbsp;</td>
<tr>
<td>microscope coverslips</td>
<td>Disposable</td>
<td>Fisher Scientific</td>
<td>12-548C</td>
<td>&nbsp;</td>
<tr>
<td>Perforated double-stick spacer tape</td>
<td>Disposable</td>
<td>Saunders Corporation (hole-punched)</td>
<td>NA</td>
<td>&nbsp;</td>
<tr>
<td>Tweezers</td>
<td>Tool</td>
<td>Various</td>
<td>NA</td>
<td>&nbsp;</td>
<tr>
<td>Brayer (roller)</td>
<td>Tool</td>
<td>Fisher Scientific</td>
<td>50820937</td>
<td>&nbsp;</td>
<tr>
<td>Chipped ice</td>
<td>Temperature Control</td>
<td>NA</td>
<td>NA</td>
<td>&nbsp;</td>
<tr>
<td>Tissues</td>
<td>Disposable</td>
<td>NA</td>
<td>NA</td>
<td>&nbsp;</td>
<tr>
<td>Calculator</td>
<td>Tool</td>
<td>Various</td>
<td>NA</td>
<td>&nbsp;</td>
<tr>
<td>Lab notebook</td>
<td>Tool</td>
<td>Various</td>
<td>NA</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>
			<div>
			* Full details for machining your own Narrow Bore Coupler are provided in Reference 3.		</div>

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.

Watch this Scientific Journal Video about Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases at JoVE.com

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

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 ...

crystallizing-membrane-proteins-for-structure-determination-using

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JoVE Journal

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29.8K Views.  Trinity College Dublin. 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.

Video: Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

The Importance of Correct Protein Concentration for Kinetics and Affinity Determination in Structure-function Analysis 

In this study, we explore the interaction between the bovine cysteine protease inhibitor cystatin B and a catalytically inactive form of papain (Fig. 1), a plant cysteine protease, by real-time label-free analysis using Biacore X100. Several cystatin B variants with point mutations in areas of interaction with papain, are produced. For each cystatin B variant we determine its specific binding concentration using calibration-free concentration analysis (CFCA) and compare the values obtained with total protein concentration as determined by A280. After that, the kinetics of each cystatin B variant binding to papain is measured using single-cycle kinetics (SCK). We show that one of the four cystatin B variants we examine is only partially active for binding. This partial activity, revealed by CFCA, translates to a significant difference in the association rate constant (ka) and affinity (KD), compared to the values calculated using total protein concentration. Using CFCA in combination with kinetic analysis in a structure-function study contributes to obtaining reliable results, and helps to make the right interpretation of the interaction mechanism.

The Importance of Correct Protein Concentration for Kinetics and Affinity Determination in Structure-function Analysis

We apply label-free protein interaction analysis using Biacore X100 for structure-function analysis of the binding of several cystatin B mutants to papain through kinetic characterization. Calibration-free concentration analysis (CFCA) measures the concentration of protein with retained binding activity without the need for a standard curve. We show that confirmation of concentrations using CFCA increases the reliability of the kinetic analysis and that kinetic constants can reliably be determined even if the activity of a recombinant protein is reduced.

In this study, we explore the interaction between the bovine cysteine protease inhibitor cystatin B and a catalytically inactive form of papain (Fig. 1), ...

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 ...

Concentration Determination of Nucleic Acids and Proteins Using the Micro-volume Bio-spec Nano Spectrophotometer

Nucleic Acid quantitation procedures have advanced significantly in the last three decades. More and more, molecular biologists require consistent small-volume analysis of nucleic acid samples for their experiments. The BioSpec-nano provides a potential solution to the problems of inaccurate, non-reproducible results, inherent in current DNA quantitation methods, via specialized optics and a sensitive PDA detector. The BioSpec-nano also has automated functionality such that mounting, measurement, and cleaning are done by the instrument, thereby eliminating tedious, repetitive, and inconsistent placement of the fiber optic element and manual cleaning.
In this study, data is presented on the quantification of DNA and protein, as well as on measurement reproducibility and accuracy. Automated sample contact and rapid scanning allows measurement in three seconds, resulting in excellent throughput. Data analysis is carried out using the built-in features of the software. The formula used for calculating DNA concentration is:
Sample Concentration = DF &middot; (OD260-OD320)&middot; NACF (1)
Where DF = sample dilution factor and NACF = nucleic acid concentration factor.
The Nucleic Acid concentration factor is set in accordance with the analyte selected1.
Protein concentration results can be expressed as &mu;g/ mL or as moles/L by entering e280 and molecular weight values respectively. When residue values for Tyr, Trp and Cysteine (S-S bond) are entered in the e280Calc tab, the extinction coefficient values are calculated as e280 = 5500 x (Trp residues) + 1490 x (Tyr residues) + 125 x (cysteine S-S bond). The e280 value is used by the software for concentration calculation.
In addition to concentration determination of nucleic acids and protein, the BioSpec-nano can be used as an ultra micro-volume spectrophotometer for many other analytes or as a standard spectrophotometer using 5 mm pathlength cells.

This communication presents data on the accuracy and reproducibility of the BioSpec-nano UV-VIS spectrophotometer for dsDNA and protein quantitation. Even with ultra-small volumes (1 to 2 L), reproducibility is excellent, while the automated wiping function improves throughput and results in minimal carryover for more precise results.

Nucleic Acid quantitation procedures have advanced significantly in the last three decades.  More and more, molecular biologists require consistent ...

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 ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...

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 ...

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, ...

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 ...