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, and especially to Jingquan Tan and Joseph Lyons, 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, P50GM073210 and U54GM094599), and FP7 COST and Marie Curie Actions (CM0902 and PIEF-GA-2009-235612).

1. Laboratory Set-up Pre-harvesting 
<ol>
 <li>In preparation for harvesting, fill the dry foam Dewar with liquid nitrogen and place it beside the microscope where harvesting is to take place. </li>
 <li>Submerge the storage puck, open end up, in the liquid nitrogen inside the foam Dewar and allow it to fully cool. </li>
 <li>Secure a micro-mount of a size that matches the crystal to be harvested on a magnetic wand (Figure 5). It is important to have on hand a number of spare magnetic wands preloaded with micro-mounts to cater for situations where it is necessary to harvest several crystals from the one bolus. Spare wands should be available at all times. </li>
 <li>Place a micropipette, tips and the precipitant solution that was used for crystal growth next to the harvesting microscope. It may be needed to cover the mesophase and to prevent drying when the well is opened. </li>
 <li>Have a notebook and pen on the bench close by and/or the computer open. These will be used to record observations regarding the quality, appearance, location, storage puck number, etc., of crystals as they are harvested, cryo-cooled and placed in storage. </li>
 <li>If an assistant is available to help with harvesting that person must understand clearly the protocol that will be followed, the order in which the various steps will take place, and their role in the overall process. </li>
</ol>
With all of the materials and equipment in place our next task is to:
2. Identify Plates and Wells that Contain Crystals
<ol>
 <li>The simplest and most direct method for finding harvestable crystals is to inspect wells by hand using a microscope with normal and with crossed polarized light. Adjusting the illuminating light intensity on the microscope can help with locating crystals. </li>
 <li>Alternatively, an imager where plates are screened automatically with normal and crossed polarized light, can be used to look for crystals. </li>
 <li>Evaluate by eye recorded digital images on a computer monitor. </li>
 <li>Clearly mark those wells with crystals for harvesting and record comments on the size, quality and location of the crystals in the mesophase in the notebook or on a computer. </li>
 <li>Remove the plate containing crystals for harvesting from the imager. </li>
</ol>
3. Opening a well with cubic mesophase. Method 1
The plates in which crystals grow by the in meso method are glass sandwich plates (Figure 2). In order to access the mesophase and the crystals therein, it is necessary to open the well. This is done by using a glass cutting tool to cut the upper coverglass that seals the well which can then be removed. 
There are several approaches for cutting and removing the coverglass from over a crystallization well. The one to use is dictated by the type of mesophase in which the crystal is found growing. This can be the very viscous and sticky cubic phase (Figure 3A) or its more fluid variant, the sponge phase (Figure 3B). In this video article we show how to open wells and to harvest crystals from both of these hosting materials. 
<ol>
 <li>Place the glass sandwich crystallization plate on the stage of a light microscope. </li>
 <li>Using a glass cutting tool score the coverglass lightly with two concentric circles situated above the spacer and just outside the perimeter of the well. With a new cutting tool, scoring requires minimal applied pressure. Replace the tool with a new one when the pressure required to score increases even fractionally; this typically occurs after opening 10 wells. </li>
 <li>Break up the glass in the space between the two scored circles for the purpose of releasing the inner coverglass. This generates lots of glass shards and dust. Clear them away with a moistened paper towel.</li>
 <li>Remove the freed coverglass by gripping it with a fine tipped tweezers and tilting it away from and off the well. In this case, the cubic phase remains stuck and in place on the baseplate of the well. </li>
 <li>Zoom in to get a clearer view of the cubic mesophase which is now ready for use in crystal harvesting. </li>
</ol>
4. Opening a Well with Cubic Mesophase. Method 2
<ol>
 <li>Using the glass cutting tool score straight parallel lines in the coverglass to one side of the well and that extend across the well itself. This enables easier tweezers access to and removal of the coverglass. </li>
</ol>
In this particular demonstration, the coverglass cracks in freeing the coverglass from the sticky spacer surface. In the process, the coverglass shifts in position and the precipitant separates from the cubic phase. When the coverglass is lifted some of the precipitant goes with it. We are left with an exposed bolus of mesophase without any surrounding precipitant. Immediately, add 1 &mu;l fresh precipitant on top of the bolus using a micropipette to prevent the mesophase from drying out and undergoing a phase change which may damage the crystals. The bolus is now ready to be used in crystal harvesting. 
5. Opening a Well with Sponge Phase. Unsuccessfully
The sponge phase is less forgiving to work with because of its ability to flow. If that flow results in the sponge phase contacting the perimeter of the well capillarity will draw out the mesophase and the crystals will be lost. An example of this happening is shown in this video clip. 
<ol>
 <li>Zoom in on the sponge phase and switch back and forth between normal and crossed polarized light to locate crystals in the sponge phase. </li>
 <li>In preparation for opening the well score and cut the coverglass as described in Section 3. In the process, the coverglass cracks. In attempting to open the well the precipitant shifts in the direction of the crack and eventually it comes in contact with the spacer. With the precipitant goes some of the sponge phase and its cargo of crystals which are lost. </li>
</ol>
In this particular sequence, the polarizers on the microscope are not completely crossed and the crystals can be seen as bright objects at the same time that the well and its contents remain visible. 
6. Opening a Well with Sponge Phase. Successfully
<ol>
 <li>Zoom in on the sponge phase and identify a crystal both in normal and crossed polarized light. </li>
 <li>Score, cut and remove a section of coverglass covering the well, as in Section 4.1. Incompletely crossed polarized light is used to keep track of the crystal. </li>
 <li>Introduce a piece of dry tissue paper through the opening in the coverglass and into the well until it just touches the precipitant solution. Wick away the solution carefully until it is almost all gone and then remove the tissue. This causes the remaining precipitant and sponge phase, with the crystal still in place, to retract under the coverglass.</li>
 <li>Score, cut and remove with a tweezers the rest of the coverglass, as in Section 4.1. In this case, the sponge phase splits; some remains in the well and some sticks to the coverglass. The crystal is in the bolus on the coverglass. Because there is very little precipitant present, the sponge phase begins to undergo a phase transition likely due to drying out. This can be seen as a ring of birefringence that migrates toward the center of the bolus. Immediately add precipitant to the bolus to halt the transition. The bolus is now ready for use in crystal harvesting. </li>
</ol>
7. Harvesting and Cryo-cooling Crystals from the Cubic Phase
<ol>
 <li>Go back and forth between normal and crossed polarized light to locate crystals in the cubic phase bolus in the open well. In this video sequence up to four birefringent crystals can be seen in the cubic phase bolus with crossed polarized light.</li>
 <li>Use a mounted cryo-loop (Figure 5) to probe the freshly exposed mesophase for crystals, to fish out crystals and then to plunge them, contained in the cryo-loop, into liquid nitrogen in the Dewar immediately upon harvesting. Ideally, the harvesting and snap-cooling should happen in one continuous and rapid motion. As little adhering mesophase as possible should be harvested with the crystal. In our experience, cryo-protectant is not needed with in meso-grown crystals. </li>
 <li>Since it is not possible to look for the crystal in the cryo-loop immediately after harvesting inspect the mesophase bolus used for harvesting to verify that the crystal is no longer there suggesting that it was harvested successfully. </li>
</ol>
8. Harvesting and Cryo-cooling Crystals from the Sponge Phase
<ol>
 <li>Go back and forth between normal and crossed polarized light to locate crystals in the sponge phase bolus in the open well. In this video sequence many birefringent crystals can be seen in the bolus under crossed polarized light. </li>
 <li>Use a mounted cryo-loop (Figure 5) to fish out crystals from the sponge phase and to plunge them, contained in or on the cryoloop, into liquid nitrogen in the Dewar immediately upon harvesting. As with harvesting from the cubic phase, ideally, the cryo-cooling process should happen immediately the crystal is harvested with as little time as possible elapsing between the actual harvesting event and plunging into liquid nitrogen. As little adhering sponge phase as possible should be harvested with the crystal. As noted, cryo-protectant is not needed with in meso-grown crystals. </li>
</ol>
9. Storing Crystals in Dewars
<ol>
 <li>Having plunged the mounted loop into liquid nitrogen place it in one of the holding slots of the storage puck in the foam Dewar. All manipulations are done with the loop, the top of the magnetic wand and the puck submerged in liquid nitrogen. </li>
 <li>Record the location and details of the harvested and cryo-cooled crystal in the notebook and/or on the computer. </li>
 <li>When the puck in the foam Dewar is full or harvesting for the day is completed transfer the puck into a shelved puck holder in a storage or a transport Dewar filled with liquid nitrogen. Crystals can be shipped in transport Dewars to the synchrotron facility for diffraction data collection. </li>
</ol>
10. Representative Results 
The objectives of the harvesting and cryo-cooling exercises demonstrated here are to transfer a crystal from the hosting mesophase into cryo-loops, to vitrify the looped crystal and to place it in storage in liquid nitrogen in a Dewar. The ideal situation is where harvesting and cryo-cooling are done in such a way that the diffraction quality of the crystal is preserved in the process. As little mesophase as possible should be harvested with the crystal. This is to make locating the crystal and centering it in the X-ray beam that much less challenging, to speed cryo-cooling with a view to vitrification, and to minimize interfering background scatter from the mesophase during diffraction data collection. Some examples of cryo-cooled samples where the crystal can and cannot be seen are shown in Figure 6. Where the crystal cannot be seen by eye it is usually necessary to resort to diffraction rastering in order to find the crystal and to center it in the beam for data collection27.
<img src="/files/ftp_upload/4001/4001fig1.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/4001/4001fig2.jpg" alt="Figure 2" /> 
Figure 2. A fully loaded and sealed 96-well glass sandwich crystallization plate. Each well contains 50 nl cubic phase and 1 &mu;l precipitant solution. For clarity, the cubic phase has been stained with Sudan Red and the precipitant solution includes Methylene Blue. From Reference 5.
<img src="/files/ftp_upload/4001/4001fig3.jpg" alt="Figure 3" /> 
Figure 3. Crystals of membrane proteins growing in the lipidic mesophase. A. The cubic phase with a crystal of bacteriorhodopsin from H. halobium. B. The sponge phase containing a crystal of the vitamin B12 receptor/transporter, BtuB, from E. coli. From Reference 25. The cubic and sponge phases have contrasting appearances as is obvious by comparing panels A and B. The cubic phase in A is extremely viscous and retains its original shape. It does not flow. This is particularly apparent at the edges of the mesophase bolus that have a roughened appearance. By contract, the sponge phase is considerably less viscous and does flow. Thus, the sponge phase fails to retain its original shape and its edges are characteristically smooth. Precipitants that include Jeffamine, PEG 400, 2-methyl-2,4-pentandiol, pentaerythritol propoxylate, butanediol and hexanediol, can result in a transition from the cubic to the sponge phase4,26.
<img src="/files/ftp_upload/4001/4001fig4.jpg" alt="Figure 4" /> 
Figure 4. The flowchart summarizes the steps involved in the production, harvesting and cryo-cooling of in meso-grown membrane protein crystals (A). Only those steps surrounded by the dashed red line, and described in detail in (B), are covered in this JoVE article. Panel A is from Reference 3. <a href="https://www.jove.com/files/ftp_upload/4001/4001fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/4001/4001fig5.jpg" alt="Figure 5" /> 
Figure 5. An empty cryo-loop mounted on a pin that is held in place on a magnetic wand. Expanded views of the pin (A) and micro-mount (B) of this important tool for harvesting and cryo-cooling are shown. The empty loop at the end of the micro-mount in B is 30 &mu;m in diameter. While having not tested other loop types extensively, we find that the MiTeGen loops shown work well with both the cubic and sponge phases. 
<img src="/files/ftp_upload/4001/4001fig6.jpg" alt="Figure 6" /> 
Figure 6. Harvested and cryo-cooled crystals of membrane proteins in cryo-loops as viewed with an in-line microscope on a synchrotron beamline. A, B. Examples of harvested crystals (caa3 cytochrome oxidase34 (A), diacylglycerol kinase, DgkA (B)) where the crystals (blue arrow) are visible through the cryo-cooled mesophase on a cryo-loop. C. Example of where the harvested crystal is not visible in the cryo-cooled mesophase on a cryo-loop. The tip of the loop is identified with a red arrow. The adhering cryo-cooled mesophase is identified with a blue arrow.

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Y., Fischer, G., Horsefield, R., Katona, G., Nyblom, M., Oberg, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Lipidic-Sponge+Phase+Screen+for+Membrane+Protein+Crystallization.">A Lipidic-Sponge Phase Screen for Membrane Protein Crystallization.</a> Structure. 16, 1003-1009 (2008).</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=Rastering+strategy+for+screening+and+centring+of+microcrystal+samples+of+human+membrane+proteins+with+a+sub-10+microm+size+X-ray+synchrotron.">Rastering strategy for screening and centring of microcrystal samples of human membrane proteins with a sub-10 microm size X-ray synchrotron.</a> 6, 587-597 (2009).</li><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=A+lipid's+eye+view+of+membrane+protein+crystallization+in+mesophases.">A lipid's eye view of membrane protein crystallization in mesophases.</a> Curr. Opin. Struct. Biol. 10, 486-497 (2000).</li><li>Briggs, J., Chung, 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+temperature-composition+phase+diagram+and+mesophase+structure+characterization+of+the+monoolein/water+system.">The temperature-composition phase diagram and mesophase structure characterization of the monoolein/water system.</a> J. Phys. Ii. 6, 723-751 (1996).</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>Misquitta, Y., Cherezov, V., Havas, F., Patterson, S., Mohan, J. M., Wells, A. J., Hart, D. J., 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=Rational+design+of+lipid+for+membrane+protein+crystallization.">Rational design of lipid for membrane protein crystallization.</a> J. Struct. Biol. 148, 169-175 (2004).</li><li>Caffrey, M., Porter, C. <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+using+Lipidic+Mesophases.">Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases.</a> J. Vis. Exp. , e1712 (2010).</li><li>Li, D., Boland, C., Walsh, K., 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=Use+of+a+robot+for+high-throughput+crystallization+of+membrane+proteins+in+lipidic+mesophase.">Use of a robot for high-throughput crystallization of membrane proteins in lipidic mesophase.</a> J. Vis. Exp. , e4000 (2012).</li><li>Lyons, J. A., Aragao, D., Slattery, O., Pisliakov, A. V., Soulimane, T., 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=Structure+insights+into+electron+transfer+in+caa(3)-type+cytochrome+oxidase.">Structure insights into electron transfer in caa(3)-type cytochrome oxidase.</a> Nature. 10, (2012).</li></ol>

No conflicts of interest declared.

In this video article we demonstrated how crystals grown in a lipidic mesophase are harvested and cryo-cooled in preparation for use in diffraction data collection and ultimately for structure determination. The hosting mesophase can be the viscous and sticky cubic phase or the more fluid sponge phase4. How the glass sandwich plates are opened and how the crystals are harvested very much depends on mesophase type. It is important therefore to know which of the two one is dealing with ahead of time. The identity of the hosting lipid and the precipitant used are important in this regard, and the physical appearance of the mesophase bolus in the crystallization well can be used to tell them apart (Figure 3). Harvesting from both mesophase types was illustrated in this article. 
Harvesting small crystals from a lipidic mesophase in glass sandwich plates is a painstaking process that requires time, skill, experience, patience and a steady hand. It is important to set aside an appropriate amount of time for harvesting and to set up the laboratory so that all the supplies and equipment are at hand in advance. A second person to assist with harvesting is not essential but is recommended. That person can help with supplying pre-marked plates to the individual doing the harvesting as well as placing cryo-cooled mounted loops with harvested crystals into the storage puck. The assistant can also play an important support role in documenting observations on the crystals made during harvesting that might prove crucial during diffraction data collection. In the absence of an assistant, a voice-activated audio-recording device could be used to advantage for documentation. 
Following the protocol described in this article will help get the Viewer up and running with crystal harvesting. However, it is important to appreciate that the process is not straightforward and that practice is needed before launching into harvesting valuable membrane protein crystals. It is recommended therefore that test plates with crystals of proteins that are not particularly valuable be experimented with first. This will provide the neophyte with valuable experience in cutting glass, removing glass shards, lifting up the coverglass from over the mesophase, using the polarizing feature on the microscope to see crystals and to track them during harvesting, and finally handling the different types of mesophases and harvesting from them. The texture of the mesophase, and by extension the ease with which crystals can be harvested, does change with time during crystallization. It is important therefore to practice harvesting with less valuable crystals but with those that have grown under the same conditions as the more valuable ones. It is possible to grow crystals of lysozyme and thaumatin by the in meso or lipidic cubic phase method28 and these should be considered by way of gaining familiarity with the materials and the method. One should also consider working with protein-free mesophase first to learn of its vagaries. 
The procedures demonstrated here were all done at a comfortable 20 &deg;C or thereabouts. It is possible to grow crystals by the in meso method at lower temperatures. Thus, monoolein as the hosting lipid in a metastable phase state can be used at 4 &deg;C 1,2,29,30. An alternative is to use the rationally designed 7.9 MAG for low temperature crystallization31. We do low temperature crysallogenesis routinely with certain membrane protein targets. In this case, crystal growth and harvesting is done in a walk-in 4 &deg;C refrigerator. Working under such conditions has its own challenges not the least of which is the need for warm and comfortable attire. 
The next step in the overall process of structure determination using macromolecular crystallography is to collect diffraction data on crystals harvested and snap-cooled as demonstrated in this article. In meso-grown crystals are usually small. However, useful diffraction data collection has been possible with crystals having a maximum dimension of 20 &mu;m9. For this purpose, micro-beam synchrotron X-radiation is used and is the focus of a separate JoVE article in this series32,33.

<table border="1">
<tbody>
 <tr>
 <td>Name of reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Components</td>
 </tr>
 <tr>
 <td>Curved tweezers</td>
 <td>Sigma</td>
 <td>F4142</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Disposable pipette tips</td>
 <td>Gilson</td>
 <td>Various</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Foam dewar</td>
 <td>Spearlab</td>
 <td>FD-500</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Glass and metal waste containers</td>
 <td>Daniels Healthcare</td>
 <td>DD479OL</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Harvesting loops</td>
 <td>MiTeGen</td>
 <td>Various</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Harvesting microscope</td>
 <td>Nikon</td>
 <td>SMZ1500</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Lab notebook</td>
 <td>Various</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Magnetic push button sample loading 			wand</td>
 <td>Hampton Research/Molecular 			Dimensions</td>
 <td>HR4-729/MD7-411</td>
 <td>Tool </td>
 </tr>
 <tr>
 <td>Original Puck (for use with 			ALS-style robots only)</td>
 <td>Crystal Positioning Systems</td>
 <td>CP-111-035</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Pipetting devices</td>
 <td>Gilson</td>
 <td>Various</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Precipitant solutions</td>
 <td>Various</td>
 <td>Various</td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Puck Bent Cryo-Tong</td>
 <td>Crystal Positioning Systems</td>
 <td>CP-111-030</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Puck Shelved Shipping Cane 			(original ALS-style) with hooked handle and locking rod </td>
 <td>Crystal Positioning Systems</td>
 <td>CP-111-029</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Purified water</td>
 <td>Millipore</td>
 <td></td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Safety goggles</td>
 <td>Various</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Sample Pin Bases - Magnetic 			(non-copper)</td>
 <td>Crystal Positioning Systems</td>
 <td>CP-111-015</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Shipping 			dewar</td>
 <td>Taylor-wharton</td>
 <td>CX100</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Tissues</td>
 <td>NA</td>
 <td>NA</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Tungsten-carbide glass cutter (TCT 			Scriber)</td>
 <td>Silverline Tools (Yeovil, UK)</td>
 <td>633657</td>
 <td>Tool</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography at JoVE.co

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

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 ...

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24.7K Views.  Trinity College Dublin. 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.

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

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 ...

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), ...

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

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

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

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

Crystallization of Membrane Proteins in Lipidic Mesophases

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

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

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

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 ...

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 ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.