There are many who contributed to this work and most are from the 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, P50GM073210 and U54GM094599), and FP7 COST Action CM0902.

1. Preparing the Crystallization Plate 
Setting up to do a crystallization trial robotically begins with the preparation of the base-plate of the glass sandwich crystallization plate (Figure 2), described in detail in Reference 2. The base-plate must first be silanized and the perforated double-stick spacer that creates the wells, must be applied to the plate. The materials and supplies needed for this are itemized under Materials. 
<ol>
 <li> Place the plate on a paper towel, apply a few drops of silanizing solution and distribute it evenly over the plate surface with a paper tissue. </li>
 <li> Remove excess silanizing solution by immersing the plate in a beaker of water. </li>
 <li> Take the plate out of the beaker, place it on a dry paper towel, silanized side up, and dry its exposed surface by wiping lightly with a paper towel. </li>
 <li> Peel off the protective cover on the perforated, double-stick spacer and apply the spacer, sticky side down, to the dry, silanized surface of the glass plate. Be sure that the spacer is positioned properly on the base-plate, with the upper left hand corner and the two adjacent sides of the spacer and the plate, coincident. </li>
 <li> Use a roller or brayer to flatten out the spacer on the base-plate and to create a tight seal between them. </li>
</ol>
Pre-assembled plates are commercially available (HR3-151, Hampton Research; MD11-55-100, Molecular Dimensions; and LCP96T-UVP70U, Swissci). For cost reasons, we prefer to assemble our own plates following the protocol above.
2. In Meso Robot 1 
The first in meso robot to be demonstrated in this video article is the one currently in use in the MS&amp;FB Group (Figure 3). Full details of the robot can be found under Materials and in the Reference section5. The materials and equipment required to set up a crystallization trial using the robot are itemized under Materials.
<ol>
 <li> Place the base-plate with spacer prepared in Section 1 on the deck of the robot. It has its own platform marked with well positions for purposes of base-plate alignment. The plate should be labeled for identification purposes. </li>
 <li> Secure the 96-well precipitant block in its own platform on the deck of the robot next to the crystallization plate. </li>
 <li> Open the block carefully by peeling off the plastic sealing cap. </li>
 <li> Turn on the humidifier and direct the stream of humid air over and across the base-plate on the deck of the robot. The humidifier has been shown to reduce evaporation of the precipitant solution and to enhance the reproducibility across the crystallization plate5. It can be turned on prior to set-up to pre-humidify the area when ambient relative humidity is low, or otherwise, just before the dispensing of mesophase commences. </li>
 <li> Initialize the robot following the manufacturer's instructions. Instrument specific instructions will not be described here in any detail because exact procedures will change over time. Essentially, what happens during initializing is that the mesophase dispensing arm on the robot is provided with reference positions in three orthogonal directions X, Y and Z. </li>
 <li> Attach the protein-laden mesophase in a Hamilton syringe, complete with dispensing needle, to Arm 1 of the robot (Figure 3). Full details of how to prepare the mesophase and how to load it into the syringe have been described in References 3,7-10. </li>
 <li> Remove the upper protective cover from the spacer on the crystallization base-plate and align the wells on the plate with fiducials on the base-plate platform. Note, for the purpose of this video, the protective cover was left in place to aid the Viewer during the alignment step. </li>
 <li> Align the needle tip of the dispensing syringe with the bottom of the uppermost well in the left hand corner of the crystallization plate (Well A1). This is done by eye adjusting the height or Z-coordinate of the dispensing arm while watching how close the needle tip comes to the bottom of the well. That the tip is just touching the bottom of the well can be judged by moving the plate from side to side on the platform whilst ever so slightly adjusting the height of the dispensing arm. </li>
 <li> Enter into the computer the Z-coordinate of the dispensing arm where the tip contacts the plate. Dispensing is programmed to take place with the needle tip 80 micrometers from this reference Z-position which corresponds to the surface of the base-plate.</li>
 <li> Position by eye the tip of the needle at the center of Well A1. Store in the controlling computer the reference X, Y coordinate of the dispensing arm where the tip is centered. This coordinate is used to define the center of the remaining wells on the plate. </li>
 <li> Prime the syringe using the appropriate command on the robot computer to ensure that fresh mesophase fills the dispensing needle and that the first and subsequent wells receive their full complement of mesophase. Wipe off extruded mesophase on the needle tip with a tissue. </li>
 <li> Start the Run Robot (X-AP) program to initiate a round of flushing steps where the precipitant dispensing tips on Arm 2 of the robot are washed and are prepared to aspirate solutions from the precipitant block.</li>
 <li> Activate the dispense function on the robot to set in motion the sequential loading of wells with protein-laden mesophase (usually 50 nl) first, a column of eight wells at a time, followed by precipitant solutions (usually 800 nl). Between loading columns, the mesophase dispensing arm returns to a parking position where the end of the needle is placed in a moistened sponge to prevent the mesophase at the tip from drying out. The process of dispensing mesophase and precipitant is repeated 11 times across the plate until the plate is filled. The loading of a 96-well plate with mesophase and precipitant takes 5 min to complete. </li>
 <li> As soon as the plate is filled and the robot arms have returned to their park positions, remove the plate from the deck of the robot and place it on a flat solid surface. </li>
 <li> Immediately, seal the plate with a cover-glass to minimize time-dependent changes in well composition. It is important to ensure that all 96 wells are fully covered, that the cover-glass is aligned square with the base-plate and that none of the cover-glass extends beyond the footprint of the base-plate. A roller or brayer is used to create a tight and a uniform seal between the cover-glass and the spacer. </li>
 <li> Inspect the plate to be sure the set up went as it should. What one is looking for is a 'fried egg on a pan' appearance in each well. The yolk of the fried egg corresponds to the mesophase. The white of the egg is the precipitant solution and the edge of the pan is the perimeter of the crystallization well. Ideally, the three are concentric. </li>
 <li> Place the plate in a temperature-controlled storage area for crystal growth. </li>
 <li> If no more plates are to be set up, remove the precipitant block from the deck of the robot, seal it tightly and put it back into storage. </li>
 <li> Remove the syringe containing the mesophase from the dispensing arm of the robot, dismantle it, and wash its parts including dispensing needle and ferrule, carefully with methanol and dry in preparation for next use. </li>
</ol>
3. In Meso Robot 2. Mosquito LCP
The Mosquito LCP robot (Figure 4) incorporates a similar mesophase dispenser to that used by Robot 1. In contrast to Robot 1, it dispenses precipitant by means of disposable tips. Programming software is used to set up the instrument for working with particular crystallization plates and precipitant blocks, and to dispense user-defined volumes of mesophase and precipitant in a specified sequence. Such details are provided by the manufacturer and will not be described here. 
<ol>
 <li> Prepare a Hamilton syringe, complete with needle and filled with protein-laden mesophase, as described above under Robot 1 and in JoVE article 17121. </li>
 <li> Clamp the syringe in position on the dispensing arm of the robot. </li>
 <li> Use the deck mounted laser sensor to establish the X and Y coordinates of the needle on the dispensing syringe. This is an important step required to ensure that the mesophase is dispensed accurately at the center of each well on the plate. </li>
 <li> With the touch of a button on the control panel, prime the syringe to expel mesophase from the needle. Remove extruded mesophase from the needle tip with a tissue in preparation for using the robot to load wells. </li>
 <li> Before placing the crystallization plate on the deck of the robot moisten the surface of the platform with a few drops of water to help hold the plate in place by capillarity. </li>
 <li> Remove the protective cover from the spacer on the glass base-plate and place it on its platform firmly to ensure good adherence. Use the three fiducials on the platform to align the 3 corner wells thereby ensuring proper positioning of the wells on the plate with respect to the dispensing needle tip. The instrument is now commanded to enter a dispensing cycle. </li>
 <li> The cycle begins with a second priming step where mesophase is dispensed onto a small glass plate immediately before moving on to load wells with fresh mesophase. The robot first loads with mesophase a vertical column of 8 wells on the plate and then uses the disposable tips to dispense precipitant solution on top of the mesophase bolus. 
 The process is repeated 11 times along the length of the plate until all wells are filled. The entire process takes about 5 min to complete. </li>
 <li> Remove the filled plate from the platform of the robot and seal it with a cover-glass as described above under Robot 1. </li>
 <li> Place the plate, properly labeled, in a temperature controlled environment for crystal growth. </li>
</ol>

4. In Meso Robot 3. Gryphon LCP
The Gryphon LCP robot (Figure 5) incorporates a similar mesophase dispensing system to that described above for Robots 1 and 2. What distinguishes it from the other two is that all 96 precipitant solutions are dispensed simultaneously. 
<ol>
 <li> Prepare the protein-laden mesophase and load it into the dispensing syringe, as described above, under Robots 1 and 2, and in JoVE article 17121. </li>
 <li> Clamp the syringe in position on the dispensing arm of the robot.</li>
 <li> Place the precipitant solution block and crystallization plate on the deck of the robot. </li>
 <li> Load the 96-tip head with precipitant solutions. A user-defined delay time is provided to facilitate aspirating viscous precipitants. </li>
 <li> Command the instrument to enter a dispensing cycle. The cycle begins with the mesophase being dispensed sequentially into the 96 wells in a process that takes about 1 min to complete. The 96-tip head then dispenses all 96 precipitant solutions at once on top of the mesophase boluses in the crystallization wells.
 The entire plate loading process is finished in 2 min. </li>
 <li> Remove the filled plate from the platform of the robot and seal it with a cover-glass as described above under Robot 1. </li>
 <li> Place the plate, properly labeled, in a temperature controlled environment for crystal growth. </li>
</ol>
5. Representative Results 
The purpose of this video article is to demonstrate how to set up automatically crystallization trials with a viscous, membrane protein-laden mesophase in glass sandwich 96-well crystallization plates. The emphasis is on automation through the use of specialized robots that require minimal manual intervention. Reproducibility and precision, in terms of volumes of mesophase and precipitant dispensed and the placement of both within the well, are expected when a robot is used. These are important from the point of view of effective screening of conditions that support crystallogenesis. They are also critical for the next step in the analysis pipeline which is to evaluate crystal growth often done with imaging equipment. Drop or bolus location is key to the imaging process, and irregular and improper placement of bolus in the well usually requires user intervention to correct for and takes time. For purposes of illustration, examples of properly and improperly loaded plates are shown in Figure 6. 
<img src="/files/ftp_upload/4000/4000fig1.jpg" alt="Figure 1" /> 
 Figure 1. The flowchart summarizes the steps involved in the production, harvesting and cryo-cooling of in meso-grown membrane protein crystals. Steps surrounded by the dashed red line are the focus of this JoVE article. Robots were used to dispense mesophase and precipitant solutions in Steps 29-32. From Reference 3. <a href="https://www.jove.com/files/ftp_upload/4000/4000fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img src="/files/ftp_upload/4000/4000fig2.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/4000/4000fig3.jpg" alt="Figure 3" /> 
 Figure 3. The in meso robot (Robot 1). The robot has two arms. Arm 1 dispenses the protein-laden lipidic mesophase. Arm 2 dispenses precipitant solutions. During filling, the 96-well plate remains fixed in position on the deck of the robot which itself is stationary. The two arms move to deliver mesophase and precipitant solutions. From Reference 3.
<img src="/files/ftp_upload/4000/4000fig4.jpg" alt="Figure 4" /> 
 Figure 4. The Mosquito LCP robot (Robot 2). In contrast to Robot 1, the deck of the Mosquito LCP, upon which sit the 96-well plate and the precipitant solution block, moves horizontally during filling. 
<img src="/files/ftp_upload/4000/4000fig5.jpg" alt="Figure 5" /> 
 Figure 5. The Gryphon LCP robot (Robot 3). During a set up, the lipidic cubic phase dispenser moves to deliver protein-laden mesophase onto the 96-well plate. The deck then translates horizontally to position the plate under the precipitant solution tip head. All 96 precipitant solutions are dispensed simultaneously. Picture sourced from the manufacturer (<a href="http://www.artrobbins.com/" target="_blank">http://www.artrobbins.com/</a>) and labeled.
<img src="/files/ftp_upload/4000/4000fig6.jpg" alt="Figure 6" /> 
 Figure 6. A fully loaded and sealed 96-well glass sandwich plate (A) and wells properly (B) and improperly (C, D) filled using the in meso robot (Robot 1). The bar code label on the plate in A is used to track the plate in the plate incubator/imaging device. In panels B-D, precipitant solution and mesophase are marked P and M, respectively. The expected 'fried egg on a pan' appearance of the well in B is characteristic of a properly loaded and sealed well. In C, the precipitant has been dispensed into the well improperly in an off-center position and has come in contact with the spacer. In C, it would appear too that the volume of mesophase is less than it should be. An example of incomplete loading with precipitant solution is shown in D. For scale, the diameter of all wells is 6 mm.

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No conflicts of interest declared.

In this video article we have demonstrated how to use a robot to automatically set up in meso crystallization trials in 96-well glass sandwich plates using a protein-laden lipidic mesophase. The robots used in this work were specially designed to include a positive displacement glass syringe for accurate and reproducible delivery of nanoliter volumes of the viscous and sticky mesophase, as originally described7.
Accuracy and precision are important features of a robot. However, these characteristics are only as good as the degree to and regularity which robot performance is evaluated and calibration is performed. It goes without saying that performance of the robot should be evaluated while individual plates are being set up. It is not appropriate to assume that the robot will function flawlessly and to leave the robot to run unattended. The attentive and observant operator should be in a position to note by a sound or an appearance when something is not working properly and to correct it immediately. Additionally, each plate should be carefully inspected by eye for uniformity of well contents as soon as the plate is sealed and before it is put away for crystallization trials. This should only require a few seconds to perform and can be done while the next plate is being loaded. Noticing, for example, that particular wells are not properly filled may highlight the fact that a certain precipitant dispensing tip is malfunctioning. Should delivery of mesophase be seen to be irregular, the relevant faulty item would need to be corrected immediately. By noting these issues and making the relevant adjustments during set up will save time and materials, including valuable lipid and membrane protein. If an imager is used to track crystal growth, accuracy and precision can also be monitored during imaging. For example, systematic problems with drop or bolus location by the imager indicate that something is not right and that corrective action somewhere upstream in the protocol is needed. 
In the interests of reliable performance therefore the robot must be calibrated on a regular schedule and as needed. Calibrations should include volume of precipitant and mesophase delivered as well as bolus and precipitant placement in the well. Obviously, wherever possible calibration should be done using volumes and materials similar to those that will be used in trials conducted in the period covered by the quality control exercise.
As important as calibration is so too is having in stock an adequate supply of robot parts and supplies. Catastrophic and unexpected failures, blockages and crashes can and do occur. In such an event, having available a replacement precipitant delivery tip, for example, might mean that an extremely valuable membrane protein preparation is used as it should and is not wasted. 
Some of the advantages of a robot include the fact that it will work essentially continuously and that it does not suffer from or 'complain of' fatigue. A robot can also be used under conditions that are not considered human-friendly, such as in the dark, under controlled lighting and environmental conditions, and at extremes of temperature. The robots demonstrated in this article were all used under ambient conditions at ~20 &deg;C. However, there are proteins and projects that require non-ambient temperatures, controlled light11,12 and an oxidizing or a reducing environment13. All of these can be catered for, with relative ease, when a crystallization robot is used.
In an earlier JoVE video article we demonstrated how crystallization trials by the in meso method are set up manually 1. The minimum volume of mesophase that can be dispensed reliably by hand is limited by the visual acuity and steady handedness of the person setting up the trials. In our experience, volumes of mesophase as low as 100 nl are readily handled. We know of one lab where the default volume dispensed manually is about 40 nl. However, considerably smaller volumes are possible using a robot. Separately, we have shown that mesophase volumes as low as 550 picoliters can be dispensed robotically14. From that work it was clear that further miniaturization by robot is possible which, if implemented, would lead to a large reduction in the amount of valuable membrane protein needed to carry out a crystallization trial. 
In this video article, three commercially available robots were used to demonstrate high-throughput membrane protein crystallization using lipidic mesophases. The first of these was developed in the MS&amp;FB Group based on our experience setting up trials manually, as described in JoVE 17121. This is the instrument we have most familiarity with and the bulk of the current article is devoted to its use. The other two robots were on demonstration in the MS&amp;FB Group at the time the article was written and the relevant footage is included here in the interests of completeness. All three robots use very much the same mesophase dispensing system, the essential feature of which is a positive displacement glass syringe5,7. They differ most significantly with regard to precipitant delivery. Robots 1 and 2 can dispense precipitant simultaneously into 8 wells, a single column on a crystallization plate at a time. In contrast, Robot 3 dispenses all 96 precipitant solutions in a single action. Robot 2 is the only instrument with disposable precipitant dispensing tips. There are pros and cons associated with the different instruments that depend on the particular application; these will not be elaborated on here. Suffice it to say that all three work and have produced crystals of membrane proteins by the in meso method.
The next steps in the overall process of structure determination by macromolecular crystallography are to harvest and to cryo-cool crystals from plates set up as described in this video article and to record and process X-ray diffraction from them. These topics are covered in separate JoVE articles in this series1,15.

<table border="1">
<tbody>
 <tr>
 <td>Name of 			reagent</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 <td>Components</td>
 </tr>
 <tr>
 <td>Brayer (roller)</td>
 <td>Fisher Scientific</td>
 <td>50820937</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Gas-tight syringes</td>
 <td>Hamilton</td>
 <td>81030</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Glass 			 coverslips</td>
 <td>Marienfeld</td>
 <td>01029990911</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Glass 			plates</td>
 <td>Marienfeld</td>
 <td>1527127092</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Gryphon LCP Robot</td>
 <td>Art Robbins</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>In meso robot</td>
 <td>Anachem/Gilson</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Lab notebook</td>
 <td>Various</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Mosquito LCP Robot</td>
 <td>TTP Labtech</td>
 <td>NA</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Perforated double-stick spacer 			tape</td>
 <td>Saunders Corporation 			(hole-punched)</td>
 <td>customized</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Precipitant solutions</td>
 <td>Various</td>
 <td>Various</td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Purified water</td>
 <td>Millipore</td>
 <td>&nbsp;NA</td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Rain-X</td>
 <td>Shell Car Care</td>
 <td>80199200</td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Syringe tips</td>
 <td>Hamilton</td>
 <td>7770-020 (gauge 22)</td>
 <td>Tool</td>
 </tr>
 <tr>
 <td>Tissues</td>
 <td>Various</td>
 <td>Various</td>
 <td>Disposable</td>
 </tr>
 <tr>
 <td>Water bottle</td>
 <td>various</td>
 <td>NA</td>
 <td>Reagent</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases at JoVE.com

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

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 ...

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17.5K Views.  Trinity College Dublin. Здесь описан робот подход к высокой пропускной способности кристаллизации мембранных белков в липидный мезофаз для использования в составе определения макромолекулярных использованием рентгеновской кристаллографии. Три роботы способны справиться с вязк?...

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

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

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

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

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

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

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

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

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

Protein Crystallization for X-ray Crystallography

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

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

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

Crystallization of Membrane Proteins in Lipidic Mesophases

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

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

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

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

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

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

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

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

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

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

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

High-throughput Purification of Affinity-tagged Recombinant Proteins

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

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

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

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

A High-throughput Method for Measurement of Glomerular Filtration Rate in Conscious Mice

The measurement of glomerular filtration rate (GFR) is the gold standard in kidney function assessment. Currently, investigators determine GFR by measuring the level of the endogenous biomarker creatinine or exogenously applied radioactive labeled inulin (3H or 14C). Creatinine has the substantial drawback that proximal tubular secretion accounts for ~50% of total renal creatinine excretion and therefore creatinine is not a reliable GFR marker. Depending on the experiment performed, inulin clearance can be determined by an intravenous single bolus injection or continuous infusion (intravenous or osmotic minipump). Both approaches require the collection of plasma or plasma and urine, respectively. Other drawbacks of radioactive labeled inulin include usage of isotopes, time consuming surgical preparation of the animals, and the requirement of a terminal experiment. Here we describe a method which uses a single bolus injection of fluorescein isothiocyanate-(FITC) labeled inulin and the measurement of its fluorescence in 1-2 &mu;l of diluted plasma. By applying a two-compartment model, with 8 blood collections per mouse, it is possible to measure GFR in up to 24 mice per day using a special work-flow protocol. This method only requires brief isoflurane anesthesia with all the blood samples being collected in a non-restrained and awake mouse. Another advantage is that it is possible to follow mice over a period of several months and treatments (i.e. doing paired experiments with dietary changes or drug applications). We hope that this technique of measuring GFR is useful to other investigators studying mouse kidney function and will replace less accurate methods of estimating kidney function, such as plasma creatinine and blood urea nitrogen.

Measurement of glomerular filtration rate (GFR) is the gold standard for kidney function assessment. Here we describe a high-throughput method which allows the determination of GFR in conscious mice by using a single bolus injection, determination of fluorescein isothiocyanate (FITC)-inulin in plasma and calculation of GFR by a two-phase exponential decay model.

The measurement of  glomerular filtration rate (GFR) is the gold standard in kidney function  assessment. Currently, investigators determine GFR by ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...