The preparation of this manuscript was partially supported by US Army Research Office (W911NF2010321) and Arizona State University startup funds to P.-L.C.

1. Teflon block preparation
<ol>
	<li>Prepare the Teflon block from chemical-resistant PTFE (polytetrafluoroethylene) resin. Make holes on the block using a general drill followed by the dimensions labeled in Figure 1.</li>
</ol>
2. Monolayer lipid preparation
NOTE: Estimated operating time: 30- 45 minutes
<ol>
	<li>Lipid stock preparation
	<ol>
		<li>Prepare a 0.01 mg/mL lipid mixture in 9:1 (v/v) chloroform/methanol using 8.91 mL of chloroform, 0.99 mL of methanol and 0.01 mL of 10 mg/mL lipids.</li>
	</ol>
	</li>
	<li>Teflon plate preparation
	<ol>
		<li>Sonicate a Teflon plate in a methanol bath for 5 minutes.</li>
		<li>Wash with hot water for 15 minutes.</li>
		<li>Rinse with distilled water.</li>
		<li>Dry the Teflon plate in a desiccator for 30 minutes and keep it under vacuum when not in use (Figure 1).</li>
	</ol>
	</li>
</ol>
3. Formation of lipid monolayer on buffer reservoir
NOTE: Estimated operating time: 1.5 hours
<ol>
	<li>Freshly prepare a lipid mixture of 0.01 mg/mL in a 9:1 mixture of chloroform/methanol on the day of the experiment.</li>
	<li>Place a Type-1 filter paper in a Petri dish and arrange the Teflon block on top of the filter paper.</li>
	<li>Fill the wells of the Teflon plate with 60 &#181;L of the buffer.</li>
	<li>Carefully add lipid mixture on top of the buffer surface drop by drop (ideally 1 &#181;L per drop) using a Hamilton syringe.</li>
	<li>Wet the filter paper with distilled water to keep humidity in the Petri dish.</li>
	<li>Incubate at room temperature for 60 minutes. The chloroform should evaporate, and a monolayer of lipid on the surface of the buffer should be formed.</li>
</ol>
4. Application of an EM grid on a lipid monolayer
NOTE: Estimated operating time: 1.5 hours
<ol>
	<li>Gently place a holey carbon-coated EM grid without glow-discharging, carbon side down, onto the top of each buffer reservoir.</li>
	<li>Carefully inject proteins into the side injection well. Use final protein concentrations in the well around 1 &#181;M of protein as a starting point. Pre-optimize the protein concentration before the experiment.</li>
	<li>Incubate for 60 minutes at room temperature.</li>
	<li>Gently inject approximately 20-30 &#181;L of buffer into the side injection port. This will raise the grid above the surface of the Teflon block.</li>
	<li>Immediately pick up the grid with a tweezer and lift it vertically off the droplet.</li>
	<li>Prepare monolayer samples for negative staining or cryo-TEM imaging (Figure 2). If the proteins form a 2D array, the image power spectrum or Fourier periodogram will show diffraction intensities at the spatial frequencies based on the crystal symmetry. If no diffraction spots are shown in the power spectrum, the proteins are unlikely to form a 2D crystal.</li>
	<li>Check the coverage of the lipid monolayer on the EM grid using a transmission electron microscope (TEM). A lower-magnified electron image is helpful to identify the morphology of the covered monolayer. Usually, the image contrast of the monolayer-covered area will be slightly lower than that of the uncovered areas. In addition, the computed 2D image power spectrum will help identify the location of the 2D crystalline.</li>
</ol>

The authors have no conflict of interest to declare.

A lipid monolayer is a powerful tool that facilitates the growth of large 2D crystals for structural studies of biological macromolecules. To successfully prepare an intact lipid monolayer at the air-water interface, it is strongly recommended that the lipids are prepared freshly on the day of the experiment, because oxidization of the lipid acyl chain could lead to packing disruption in the monolayer and adversely affect the resulting crystal formation. Purchased lipids in powder form should be dissolved using a mixture of chloroform:methanol solvent, which is commonly used to dissolve phospholipids35. The chloroform:methanol solvent can be premade and stored in a sealed glass vial. While the dissolved lipids are applied at the air-water interface, wait until the chloroform:methanol solvent completely evaporates from the buffer. Proper waiting time for the formation of lipid monolayer and the formation of 2D crystals is essential in this experiment. Typically, the formation of a lipid monolayer and 2D arrays can take place within approximately 30 to 60 minutes of incubation time18.
When a lipid monolayer is properly formed, ensure that the 2D crystal array formation can be promoted by the lipid effectively. The buffer composition should be optimized to stabilize the establishment of 2D arrays, especially when DOGS-NTA-Ni lipid is used18,36. The success of the lipid monolayer crystallization for this lipid may be affected if there are high concentrations of EDTA (ethylenediaminetetraacetate), DTT (1,4-dithiothreitol), or various ions in the buffer. The soluble chelator EDTA can disrupt the lipid-metal ion-protein complex36. One example of this is the streptavidin protein interaction with iminodiacetate-Cu (II) lipid; the addition of EDTA removes Cu (II) and interrupts the lipid-metal ion-protein complex20. Hence, the sample may need to be diluted or buffer exchanged to remove EDTA, allowing a stronger binding of protein and promoting crystal formation. The optimization of the buffer needs to be done before it is added to the buffer reservoir in the Teflon block.
To preserve the high-resolution structural detail of the 2D crystal, maintaining the specimen flatness is critical. However, due to surface tension, differential heat capacities between the specimen and support or human intervention, it is not trivial to maintain the flatness of the 2D crystal during the sample transfer. When the EM grid is lifted from the buffer reservoir, one must operate it with as little disturbance as possible to leave the monolayer interface intact. Any drastic rotation or movement of the grid may fracture the monolayer interface. This also applies for sample freezing steps, where the grid with crystals must be handled with care and rapidly frozen to avoid ice crystals, which can damage the newly formed 2D protein crystals.
One may use a lacey carbon-coated grid to support monolayers in different sizes, depending on the hole sizes on the lacey carbon film. The flatness of the monolayer interface may be affected by various hole sizes on the lacey carbon film, which may help screen a flat monolayer with the diffraction intensities in isotropic directions.
There are four common methods used to grow and develop 2D crystals for structural studies: dialysis, dilution, hydrophobic adsorption and lipid monolayer37,38,39. The dialysis method removes the detergents that are used to solubilize membrane proteins in the early step of purification. The 2D crystals are grown by reconstituting detergent-solubilized membrane proteins into lipid bilayers under favorable conditions35. For the reconstitution to take place, detergents need to be removed by dialysis. Therefore, the dialysis method helps to drive the formation of 2D crystals40. However, the dialysis method is usually time-consuming, especially for detergents with a small critical micelle concentration (CMC). The formation of 2D arrays can also be achieved via a controlled dilution. This method provides a set ratio for the protein-lipid-detergent mixture to induce crystal formation if the detergent concentration drops below its CMC. This could be facilitated by a specific dilution machine to carry out the work starting with a higher initial protein concentration41. In addition, hydrophobic adsorption prompts the growth of 2D crystals by using polystyrene beads (e.g., Bio-beads) to remove the low CMC detergents42. If the protein of interest is solubilized with high CMC detergents, this method is not ideal. In contrast, the lipid monolayer method is versatile and provides non-convoluted steps for 2D crystallization43. As discussed here, it requires a simple apparatus and some materials to conduct the experiments. With our method, the total time needed for the experiment can be shortened to 4 hours17. Even though pre-optimization of certain parameters, such as protein concentration and buffer components, may still be required beforehand, it is possible to maximize one&#39;s efficiency in routine lab work.
For crystallizing integral membrane proteins using a lipid monolayer, one should test whether the detergent concentration for solubilizing protein and lipids disturbs the monolayer interface. Generally, it is suggested to use the detergent concentration lower than the CMC 44. In cases that the monolayer interface is sensitive to the detergent existence, an alternative approach can be to form the lipid monolayer using fluoro-lipids, which are hardly solubilized by detergent and expand the applicability of integral membrane protein crystallization using the monolayer method45.
Although the lipid monolayer method is simple and less costly, the reproducibility of the monolayer preparation depends on various factors, such as ambient humidity and temperature. The success of the sample transfer for cryo-EM imaging relies on careful specimen handling. In some cases, it may be challenging to obtain high-resolution results by a novice or in the beginning phase of a new research project.
In conclusion, the lipid monolayer method offers an opportunity for studying protein structures due to its simplicity. It may provide an alternative approach to facilitate the process of 2D crystal formation.

Electron diffraction through 2D crystals or helical arrays of proteins can achieve sub-nanometer resolutions in favorable cases1,2,3. Of particular interest are reconstituted 2D membrane protein arrays or crystals in their near-native environments1. Because a crystal acts as a signal amplifier enhancing the intensities of the structural factors at specific spatial frequencies, electron crystallography allows probing a target with a smaller size at high resolutions, such as small molecules, than those for single-particle cryo-EM. The electron beam can be diffracted by an ordered 2D array of proteins, generating a diffraction pattern or a lattice image depending on where the image plane is recorded on the detector4. The diffracted intensities can then be extracted and processed to reconstruct a 2D projection structure of the crystal. Electrons have a larger scattering cross-section than X-rays and its scattering mostly follows the Rutherford model based on the Coulomb interaction between the electrons and the charged atoms in the molecule5. The thicknesses of 2D membrane crystals are usually less than 100 nm, suitable for electron transmission without dynamical scattering occurring within specimen6. Electron crystallographic studies have been shown to be a powerful tool to probe high-resolution structural information of membrane proteins and lipid-protein interactions7,8,9,10,11,12,13,14,15,16,17.
A lipid monolayer is one single lipid layer composed of phospholipids densely packed at an air-water interface6, which can assist the 2D array formation for soluble proteins or peripheral membrane proteins18. Depending on the density of the lipids and their lateral pressure, the lipid molecules can form an ordered 2D array on the air-water interface with their acyl chains extended to the air and hydrophilic headgroups exposed in the aqueous solution1,6,19. The lipid headgroup can interact with proteins via electrostatic interaction or can be modified to provide an affinity tag to bind a specific protein domain. For example, the DOGS-NTA-Ni (1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl]2- Ni2+) is often used in forming a lipid monolayer to bind the proteins with a poly-histidine tag20,21,22. Also, the cholera toxin B can bind a particular pentasaccharide of ganglioside GM1 in a lipid monolayer for structural studies23,24. By anchoring the proteins on the lipid headgroups, the lipid monolayer can assist the formation of the 2D arrays that are thin for high-resolution electron crystallographic studies. The lipid monolayer technique has been used in electron crystallography for structural studies of proteins, such as streptavidin2,25, annexin V26, cholera toxin27, E. coli gyrase B subunit28, E. coli RNA polymerase25,29,30, carboxysome shell proteins31 and the capsid proteins of the HIV-132 and Moloney murine leukemia virus33. Due to the stability and chemical property of the lipid monolayer, different applications for sample preparation have been explored for cryo-EM imaging34. However, optimization will be needed for protein array formation.
Here, we provide extensive details of the general preparation of lipid monolayers for cryo-EM imaging and some considerations that could affect the quality of the formed monolayers.

<table><tbody><tr><td>14:0 PC (DMPC)</td><td>Avanti Lipids</td><td>850345</td><td>1,2-dimyristoyl-&lt;em&gt;sn&lt;/em&gt;-glycero-3-phosphocholine,&lt;br/&gt; 1 x 25 mg, 10 mg/mL, 2.5 mL</td></tr><tr><td>Bulb for small pipets</td><td>Fisher Scientific</td><td>03-448-21</td><td></td></tr><tr><td>Chloroform</td><td>Sigma-Aldrich</td><td>C2432</td><td></td></tr><tr><td>Desiccator vacuum</td><td>Southern Labware</td><td>55207</td><td></td></tr><tr><td>EM grids</td><td>Electron Microscopy Sciences</td><td>CF413-50</td><td>CF-1.2/1.3-4C 1.2 &amp;micro;m hole, 1.3 &amp;micro;m space</td></tr><tr><td>Filter paper</td><td>GE Healthcare Life Sciences</td><td>1001-090</td><td>Diameter 90 mm</td></tr><tr><td>Glass Pasteur pipets</td><td>Fisher Scientific</td><td>13-678-20A</td><td></td></tr><tr><td>Hamilton syringe (25 &amp;micro;L)</td><td>Hamilton Company</td><td>80465</td><td></td></tr><tr><td>Hamilton syringe (250 &amp;micro;L)</td><td>Hamilton Company</td><td>81165</td><td></td></tr><tr><td>Hamilton syringe (5 &amp;micro;L)</td><td>Hamilton Company</td><td>87930</td><td></td></tr><tr><td>Hamilton syringe (500 &amp;micro;L)</td><td>Hamilton Company</td><td>203080</td><td></td></tr><tr><td>Methanol</td><td>Sigma-Aldrich</td><td>M1775-1GA</td><td></td></tr><tr><td>Petri dish</td><td>VWR</td><td>25384-342</td><td>100 mm &amp;times; 15 mm</td></tr><tr><td>Teflon block</td><td>Grainger</td><td>55UK05</td><td>60 &amp;micro;L wells with side injection ports, manually made</td></tr><tr><td>Tweezers</td><td>Electron Microscopy Sciences</td><td>78325</td><td>Various styles</td></tr><tr><td>Ultra-pure water</td><td></td><td></td><td></td></tr><tr><td>Ultrasonic cleaner</td><td>VWR</td><td>97043-996</td><td></td></tr></tbody></table>

sample preparation using a lipid monolayer method for electron crystallographic studies

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown into highly ordered two-dimensional (2D) crystals under favorable conditions. The quality of the grown 2D crystals is crucial to the resolution of the final reconstruction via 2D image processing. Over the years, lipid monolayers have been used as a supporting layer to foster the 2D crystallization of peripheral membrane proteins as well as soluble proteins. This method can also be applied to 2D crystallization of integral membrane proteins but requires more extensive empirical investigation to determine detergent and dialysis conditions to promote partitioning to the monolayer. A lipid monolayer forms at the air-water interface such that the polar lipid head groups remain hydrated in the aqueous phase and the non-polar, acyl chains, tails partition into the air, breaking the surface tension and flattening the water surface. The charged nature or distinctive chemical moieties of the head groups provide affinity for proteins in solution, promoting binding for 2D array formation. A newly formed monolayer with the 2D array can be readily transfer into an electron microscope (EM) on a carbon-coated copper grid used to lift and support the crystalline array. In this work, we describe a lipid monolayer methodology for cryogenic electron microscopic (cryo-EM) imaging.

A lipid monolayer deposited on the EM grid can be visualized under a transmission electron microscope (TEM) without staining. The monolayer presence can be recognized by the contrast difference from the area without any specimen in the beam path. Areas that have lipid monolayer coverage have lower local contrast than the ones with no coverage, since the electron beam through the empty holes has no scattering and shows a brighter illumination (Figure 3).
To screen the conditions for 2D monolayer crystallization, investigate the image of the specimen using negative-stain EM or cryo-EM imaging. Whether the monolayer covers the holes could be observed by a stronger edge contrast when the stage is tilted. If a crystal is formed, the image power spectrum or periodogram will show the Fourier or diffraction spots. Crystals in small sizes (&#60; 200 nm) and randomly oriented may show high-intensity rings in the power spectrum. If the crystals are not grown, the power spectrum will not show Fourier spots.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63015/63015fig01.jpg" /> 
Figure 1. Teflon block used for lipid monolayer formation. A Teflon block is shown in top (above) and cross-section (below) views. It contains 12 wells arranged in six rows and two columns. The main buffer well labeled A, shown in darker blue, in the middle is 4 mm in diameter and 5 mm deep with 60 &#181;L sample volumes. The monolayer is formed on top of the buffer reservoir in the main well and proteins are injected through the side port labeled B shown in light blue. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63015/63015fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63015/63015fig02.jpg" /> 
Figure 2. Schematic presentation of the development of 2D crystal on lipid monolayer. The buffer reservoir is first filled with the buffer, followed by the lipid deposited on top of the well. The carbon side of an EM grid is placed on the lipid monolayer, which will interact with the hydrophobic tails. First, proteins are injected through the side injection well B (step 1). Over time, the proteins are arranged into a 2D crystal lattice and interact with the lipid head groups (step 2). To pick up the EM grid for further analysis, a tweezer is used to lift the grid off the buffer reservoir vertically (step 3). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63015/63015fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63015/63015fig03.jpg" /> 
Figure 3. Example of the lipid monolayer coverage on an EM grid. A monolayer- and holey carbon-coated EM grid without staining was imaged using a Philips CM12 transmission electron microscope at room temperature. The orange arrow indicates the area that has no lipid monolayer coverage, and the blue arrows indicate the full coverage of the lipid monolayer in the grid hole. Scale bar is 0.5 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63015/63015fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies at JoVE.com

Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Lipid monolayers have been used as a foundation for forming two-dimensional (2D) protein crystals for structural studies for decades. They are stable at the air-water interface and can serve as a thin supporting material for electron imaging. Here we present the proven steps on preparing lipid monolayers for biological studies.

Electron crystallography is a powerful tool for high-resolution structure determination. Macromolecules such as soluble or membrane proteins can be grown ...

sample-preparation-using-lipid-monolayer-method-for-electron

Nanyang Technological University

Research

JoVE Journal

Biochemistry

3.9K Views.  Arizona State University. Lipid monolayers have been used as a foundation for forming two-dimensional (2D) protein crystals for structural studies for decades. They are stable at the air-water interface and can serve as a thin supporting material for electron imaging. Here we present the proven steps on preparing lipid monolayers for biological studies.

Video: Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Preparation and Delivery of Protein Microcrystals in Lipidic Cubic Phase for Serial Femtosecond Crystallography

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural information, typically obtained by crystallography; however MPs are difficult to crystallize. Recent progress in MP structural determination has benefited greatly from the development of lipidic cubic phase (LCP) crystallization methods, which typically yield well-diffracting, but often small crystals that suffer from radiation damage during traditional crystallographic data collection at synchrotron sources. The development of new-generation X-ray free-electron laser (XFEL) sources that produce extremely bright femtosecond pulses has enabled room temperature data collection from microcrystals with no or negligible radiation damage. Our recent efforts in combining LCP technology with serial femtosecond crystallography (LCP-SFX) have resulted in high-resolution structures of several human G protein-coupled receptors, which represent a notoriously difficult target for structure determination. In the LCP-SFX technique, LCP is recruited as a matrix for both growth and delivery of MP microcrystals to the intersection of the injector stream with an XFEL beam for crystallographic data collection. It has been demonstrated that LCP-SFX can substantially improve the diffraction resolution when only sub-10 &#181;m crystals are available, or when the use of smaller crystals at room temperature can overcome various problems associated with larger cryocooled crystals, such as accumulation of defects, high mosaicity and cryocooling artifacts. Future advancements in X-ray sources and detector technologies should make serial crystallography highly attractive and practicable for implementation not only at XFELs, but also at more accessible synchrotron beamlines. Here we present detailed visual protocols for the preparation, characterization and delivery of microcrystals in LCP for serial crystallography experiments. These protocols include methods for conducting crystallization experiments in syringes, detecting and characterizing the crystal samples, optimizing crystal density, loading microcrystal laden LCP into the injector device and delivering the sample to the beam for data collection.

We describe procedures for the preparation and delivery of membrane protein microcrystals in lipidic cubic phase for serial crystallography at X-ray free-electron lasers and synchrotron sources. These protocols can also be applied for incorporation and delivery of soluble protein microcrystals, leading to substantially reduced sample consumption compared to liquid injection.

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural ...

From Constructs to Crystals &#8211; Towards Structure Determination of &#946;-barrel Outer Membrane Proteins

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% percent of known structures. There are two types of membrane proteins, &#945;-helical and &#946;-barrel. While &#945;-helical membrane proteins can be found in nearly all cellular membranes, &#946;-barrel membrane proteins can only be found in the outer membranes of mitochondria, chloroplasts, and Gram-negative bacteria. One common bottleneck in structural studies of membrane proteins in general is getting enough pure sample for analysis. In hopes of assisting those interested in solving the structure of their favorite &#946;-barrel outer membrane protein (OMP), general protocols are presented for the production of target &#946;-barrel OMPs at levels useful for structure determination by either X-ray crystallography and/or NMR spectroscopy. Here, we outline construct design for both native expression and for expression into inclusion bodies, purification using an affinity tag, and crystallization using detergent screening, bicelle, and lipidic cubic phase techniques. These protocols have been tested and found to work for most OMPs from Gram-negative bacteria; however, there are some targets, particularly for mitochondria and chloroplasts that may require other methods for expression and purification. As such, the methods here should be applicable for most projects that involve OMPs from Gram-negative bacteria, yet the expression levels and amount of purified sample will vary depending on the target OMP.

&#946;-barrel outer membrane proteins (OMPs) serve many functions within the outer membranes of Gram-negative bacteria, mitochondria, and chloroplasts. Here, we hope to alleviate a known bottleneck in structural studies by presenting protocols for the production of &#946;-barrel OMPs in sufficient quantities for structure determination by X-ray crystallography or NMR spectroscopy.

Membrane proteins serve important functions in cells such as nutrient transport, motility, signaling, survival and virulence, yet constitute only ~1% ...

Chemistry

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

Bioengineering

Improving High Viscosity Extrusion of Microcrystals for Time-resolved Serial Femtosecond Crystallography at X-ray Lasers

High-viscosity micro-extrusion injectors have dramatically reduced sample consumption in serial femtosecond crystallographic experiments (SFX) at X-ray free electron lasers (XFELs). A series of experiments using the light-driven proton pump bacteriorhodopsin have further established these injectors as a preferred option to deliver crystals for time-resolved serial femtosecond crystallography (TR-SFX) to resolve structural changes of proteins after photoactivation. To obtain multiple structural snapshots of high quality, it is essential to collect large amounts of data and ensure clearance of crystals between every pump laser pulse. Here, we describe in detail how we optimized the extrusion of bacteriorhodopsin microcrystals for our recent TR-SFX experiments at the Linac Coherent Light Source (LCLS). The goal of the method is to optimize extrusion for a stable and continuous flow while maintaining a high density of crystals to increase the rate at which data can be collected in a TR-SFX experiment. We achieve this goal by preparing lipidic cubic phase with a homogenous distribution of crystals using a novel three-way syringe coupling device followed by adjusting the sample composition based on measurements of the extrusion stability taken with a high-speed camera setup. The methodology can be adapted to optimize the flow of other microcrystals. The setup will be available for users of the new Swiss Free Electron Laser facility.

The success of a time-resolved serial femtosecond crystallography experiment is dependent on efficient sample delivery. Here, we describe protocols to optimize the extrusion of bacteriorhodopsin microcrystals from a high viscosity micro-extrusion injector. The methodology relies on sample homogenization with a novel three-way coupler and visualization with a high-speed camera.

High-viscosity micro-extrusion injectors have dramatically reduced sample consumption in serial femtosecond crystallographic experiments (SFX) at X-ray ...

A Sample Preparation Pipeline for Microcrystals at the VMXm Beamline

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality have been seen for microcrystals with the development of beamline optics, beam stability and variable beam size focusing from submicron to microns, such as at the VMXm beamline at Diamond Light Source1. Further improvements in data quality will be gained through improvements in sample environment and sample preparation. Microcrystals inherently generate weaker diffraction, therefore improving the signal-to-noise is key to collecting quality X-ray diffraction data and will predominantly come from reductions in background noise. Major sources of X-ray background noise in a diffraction experiment are from their interaction with the air path before and after the sample, excess crystallization solution surrounding the sample, the presence of crystalline ice and scatter from any other beamline instrumentation or X-ray windows. The VMXm beamline comprises instrumentation and a sample preparation protocol to reduce all these sources of noise.
Firstly, an in-vacuum sample environment at VMXm removes the air path between X-ray source and sample. Next, sample preparation protocols for macromolecular crystallography at VMXm utilize a number of processes and tools adapted from cryoTEM. These include copper grids with holey carbon support films, automated blotting and plunge cooling robotics making use of liquid ethane. These tools enable the preparation of hundreds of microcrystals on a single cryoTEM grid with minimal surrounding liquid on a low-noise support. They also minimize the formation of crystalline ice from any remaining liquid surrounding the crystals. 
We present the process for preparing and assessing the quality of soluble protein microcrystals using visible light and scanning electron microscopy before mounting the samples on the VMXm beamline for X-ray diffraction experiments. We will also provide examples of good quality samples as well as those which require further optimization and strategies to do so.

The signal-to-noise ratio of data is one of the most important considerations in performing X-ray diffraction measurements from microcrystals. The VMXm beamline provides a low-noise environment and microbeam for such experiments. Here, we describe sample preparation methods for mounting and cooling microcrystals for VMXm and other microfocus macromolecular crystallography beamlines.

The mounting of microcrystals (&#60;10 &#181;m) for single crystal cryo-crystallography presents a non-trivial challenge. Improvements in data quality ...

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 ...

Biology

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 ...

The Peel-Blot Technique: A Cryo-EM Sample Preparation Method to Separate Single Layers From Multi-Layered or Concentrated Biological Samples

The peel-blot cryo-EM grid preparation technique is a significantly modified back-injection method with the objective of achieving a reduction in layers of multi-layered samples. This removal of layers prior to plunge freezing can aid in reducing sample thickness to a level suitable for cryo-EM data collection, improving sample flatness, and facilitating image processing. The peel-blot technique allows for the separation of multilamellar membranes into single layers, of layered 2D crystals into individual crystals, and of stacked, sheet-like structures of soluble proteins to likewise be separated into single layers. The high sample thickness of these types of samples frequently poses insurmountable problems for cryo-EM data collection and cryo-EM image processing, especially when the microscope stage must be tilted for data collection. Furthermore, grids of high concentrations of any of these samples can be prepared for efficient data collection since sample concentration prior to grid preparation can be increased and the peel-blot technique adjusted to result in a dense distribution of single-layered specimen.

The peel-blot technique is a cryo-EM grid preparation method that allows for the separation of multilayered and concentrated biological samples into single layers to reduce thickness, increase sample concentration, and facilitate image processing.

The peel-blot cryo-EM grid preparation technique is a significantly modified back-injection method with the objective of achieving a reduction in layers ...

Fabrication of Streptavidin Affinity Grids for Cryo-EM Structural Studies

Streptavidin-Affinity Grid Fabrication for Cryo-Electron Microscopy Sample Preparation

Streptavidin affinity grids provide strategies to overcome many commonly encountered cryo-electron microscopy (cryo-EM) sample preparation challenges, including sample denaturation and preferential orientations that can occur due to the air-water interface. Streptavidin affinity grids, however, are currently utilized by few cryo-EM labs because they are not commercially available and require a careful fabrication process. Two-dimensional streptavidin crystals are grown onto a biotinylated lipid monolayer that is applied directly to standard holey-carbon cryo-EM grids. The high-affinity interaction between streptavidin and biotin allows for the subsequent binding of biotinylated samples that are protected from the air-water interface during cryo-EM sample preparation. Additionally, these grids provide a strategy for concentrating samples available in limited quantities and purifying protein complexes of interest directly on the grids. Here, a step-by-step, optimized protocol is provided for the robust fabrication of streptavidin affinity grids for use in cryo-EM and negative-stain experiments. Additionally, a trouble-shooting guide is included for commonly experienced challenges to make the use of streptavidin affinity grids more accessible to the larger cryo-EM community.

Author Spotlight: Enhancing Cryo-EM Sample Preparation with Streptavidin-Biotin Approach

A step-by-step protocol for fabricating streptavidin affinity grids is provided for use in structural studies of challenging macromolecular samples by cryo-electron microscopy.

Streptavidin affinity grids provide strategies to overcome many commonly encountered cryo-electron microscopy (cryo-EM) sample preparation challenges, ...

Assembly of Cell Mimicking Supported and Suspended Lipid Bilayer Models for the Study of Molecular Interactions

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial protective barrier for all cell types, separating the internal cellular components from the extracellular environment. These membranes are composed largely of a lipid bilayer, which contains outer hydrophilic head groups and inner hydrophobic tail groups, along with various proteins and cholesterol. The composition and structure of the lipids themselves play a crucial role in regulating biological function, including interactions between cells and the cellular microenvironment, which may contain pharmaceuticals, biological toxins, and environmental toxicants. In this study, methods to formulate uni-lipid and multi-lipid supported and suspended cell mimicking lipid bilayers are described. Previously, uni-lipid phosphatidylcholine (PC) lipid bilayers as well as multi-lipid placental trophoblast-inspired lipid bilayers were developed for use in understanding molecular interactions. Here, methods for achieving both types of bilayer models will be presented. For cell mimicking multi-lipid bilayers, the desired lipid composition is first determined via lipid extraction from primary cells or cell lines followed by liquid chromatography-mass spectrometry (LC-MS). Using this composition, lipid vesicles are fabricated using a thin-film hydration and extrusion method and their hydrodynamic diameter and zeta potential are characterized. Supported and suspended lipid bilayers can then be formed using quartz crystal microbalance with dissipation monitoring (QCM-D) and on a porous membrane for use in a parallel artificial membrane permeability assay (PAMPA), respectively. The representative results highlight the reproducibility and versatility of in vitro cell membrane lipid bilayer models. The methods presented can aid in rapid, facile assessment of the interaction mechanisms, such as permeation, adsorption, and embedment, of various molecules and macromolecules with a cell membrane, helping in the screening of drug candidates and prediction of potential cellular toxicity.

This protocol describes the formation of cell mimicking uni-lipid and multi-lipid vesicles, supported lipid bilayers, and suspended lipid bilayers. These in vitro models can be adapted to incorporate a variety of lipid types and can be used to investigate various molecule and macromolecule interactions.

Model cell membranes are a useful screening tool with applications ranging from early drug discovery to toxicity studies. The cell membrane is a crucial ...