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...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>14:0 PC (DMPC)</td>
			<td>Avanti Lipids</td>
			<td>850345</td>
			<td>1,2-dimyristoyl-sn-glycero-3-phosphocholine, 
			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 &#181;m hole, 1.3 &#181;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 &#181;L)</td>
			<td>Hamilton Company</td>
			<td>80465</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton syringe (250 &#181;L)</td>
			<td>Hamilton Company</td>
			<td>81165</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton syringe (5 &#181;L)</td>
			<td>Hamilton Company</td>
			<td>87930</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton syringe (500 &#181;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 &#215; 15 mm</td>
		</tr>
		<tr>
			<td>Teflon block</td>
			<td>Grainger</td>
			<td>55UK05</td>
			<td>60 &#181;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 ...

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

Universidad de Cartagena UDEC

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Sample Preparation for Mass Spectrometry-based Identification of RNA-binding Regions

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most cases, the action of noncoding RNAs is mediated by proteins whose functions are in turn regulated by these interactions with noncoding RNAs. Consistent with this, a growing number of proteins involved in nuclear functions have been reported to bind RNA and in a few cases the RNA-binding regions of these proteins have been mapped, often through laborious, candidate-based methods.
Here, we report a detailed protocol to perform a high-throughput, proteome-wide unbiased identification of RNA-binding proteins and their RNA-binding regions. The methodology relies on the incorporation of a photoreactive uridine analog in the cellular RNA, followed by UV-mediated protein-RNA crosslinking, and mass spectrometry analyses to reveal RNA-crosslinked peptides within the proteome. Although we describe the procedure for mouse embryonic stem cells, the protocol should be easily adapted to a variety of cultured cells.

We describe a protocol to identify RNA-binding proteins and map their RNA-binding regions in live cells using UV-mediated photocrosslinking and mass spectrometry.

Noncoding RNAs play important roles in several nuclear processes, including regulating gene expression, chromatin structure, and DNA repair. In most ...

A Plasma Sample Preparation for Mass Spectrometry using an Automated Workstation

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves numerous incubation and liquid transfer steps in order to achieve denaturation, reduction, alkylation, and cleavage. Adapting this workflow onto an automated workstation can increase efficiency and reduce coefficients of variance, thereby providing more reliable data for statistical comparisons between sample types. We previously described an automated proteomic sample preparation workflow1. Here, we report the development of a more efficient and better controlled workflow with the following advantages: 1) The number of liquid transfer steps is reduced from nine to six by combining reagents; 2) Pipetting time is reduced by selective tip pipetting using a 96-position pipetting head with multiple channels; 3) Potential throughput is increased by the availability of up to 45 deck positions; 4) Complete enclosure of the system provides improved temperature and environmental control and reduces the potential for contamination of samples or reagents; and 5) The addition of stable isotope labeled peptides, as well as &#946;-galactosidase protein, to each sample makes monitoring and quality control possible throughout the entire process. These hardware and process improvements provide good reproducibility and improve intra-assay and inter-assay precision (CV of less than 20%) for LC-MS based protein and peptide quantification. The entire workflow for digesting 96 samples in a 96-well plate can be completed in approximately 5 hours.

Here, we present an automated plasma protein digestion method for mass spectrometry-based quantitative proteomic analysis. In this protocol, the liquid transferring and incubation steps for protein denaturation, reduction, alkylation, and trypsin digestion reactions are streamlined and automated. It takes approximately five hours to prepare a 96-well plate with desired precision.

Sample preparation for mass spectrometry analysis in proteomics requires enzymatic cleavage of proteins into a peptide mixture. This process involves ...

Sample Preparation for Probe Electrospray Ionization Mass Spectrometry

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ratios (m/z), which are useful to deduce molecular weights and structures. While it is essentially a destructive analytical method, recent advancements in the ambient ionization technique have enabled us to acquire data while leaving tissue in a relatively intact state in terms of integrity. Probe electrospray ionization (PESI) is a so-called direct method because it does not require complex and time-consuming pretreatment of samples. A fine needle serves as a sample picker, as well as an ionization emitter. Based on the very sharp and fine property of the probe tip, destruction of the samples is minimal, allowing us to acquire the real-time molecular information from living things in situ. Herein, we introduce three applications of PESI-MS technique that will be useful for biomedical research and development. One involves the application to solid tissue, which is the basic application of this technique for the medical diagnosis. As this technique requires only 10 mg of the sample, it may be very useful in the routine clinical settings. The second application is for in vitro medical diagnostics where human blood serum is measured. The ability to measure fluid samples is also valuable in various biological experiments where a sufficient volume of sample for conventional analytical techniques cannot be provided. The third application leans toward the direct application of probe needles in living animals, where we can obtain real-time dynamics of metabolites or drugs in specific organs. In each application, we can infer the molecules that have been detected by MS or use artificial intelligence to obtain a medical diagnosis.

This article introduces sample preparation methods for a unique real-time analytical method based on the ambient mass spectrometry. This method lets us perform real-time analysis of the biological molecules in vivo without any special pretreatments.

Mass spectrometry (MS) is a powerful tool in analytical chemistry because it provides very accurate information about molecules, such as mass-to-charge ...

TMT Sample Preparation for Proteomics Facility Submission and Subsequent Data Analysis

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view of the impact of a disease, treatment, or other condition on the proteome as a whole. This report provides a detailed protocol for the extraction, quantification, precipitation, digestion, labeling, and subsequent data analysis of protein samples. Our optimized TMT labeling protocol requires a lower tag-label concentration and achieves consistently reliable data. We have used this protocol to evaluate protein expression profiles in a variety of mouse tissues (i.e., heart, skeletal muscle, and brain) as well as cells cultured in vitro. In addition, we demonstrate how to evaluate thousands of proteins from the resulting dataset.

We present an optimized tandem mass tag (TMT) labeling protocol that includes detailed information for each of the following steps: protein extraction, quantification, precipitation, digestion, labeling, submission to a proteomics facility, and data analyses.

Proteomic technologies are powerful methodologies that can aid our understanding of mechanisms of action in biological systems by providing a global view ...

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures and information on more challenging systems. Sample preparation for cryo-EM is undergoing a similar revolution with new approaches being developed to supersede the traditional blotting systems. These include the use of piezo-electric dispensers, pin printing and direct spraying. As a result of these developments, the speed of grid preparation is going from seconds to milliseconds, providing new opportunities, especially in the field of time-resolved cryo-EM where proteins and substrates can be rapidly mixed before plunge freezing, trapping short lived intermediate states. Here we describe, in detail, a standard protocol for making grids on our in-house time-resolved EM device both for standard fast grid preparation and also for time-resolved experiments. The protocol requires a minimum of about 50 &#181;L sample at concentrations of &#8805; 2 mg/mL for the preparation of 4 grids. The delay between sample application and freezing can be as low as 10 ms. One limitation is increased ice thickness at faster speeds and compared to the blotting method. We hope this protocol will aid others in designing their own grid making devices and those interested in designing time-resolved experiments.

Here, we provide a detailed protocol for the use of a rapid grid making device for both fast grid-making and for rapid mixing and freezing to conduct time-resolved experiments.

The field of cryo-electron microscopy (cryo-EM) is rapidly developing with new hardware and processing algorithms, producing higher resolution structures ...

Workflow and Tools for Crystallographic Fragment Screening at the Helmholtz-Zentrum Berlin

Fragment screening is a technique that helps to identify promising starting points for ligand design. Given that crystals of the target protein are available and display reproducibly high-resolution X-ray diffraction properties, crystallography is among the most preferred methods for fragment screening because of its sensitivity. Additionally, it is the only method providing detailed 3D information of the binding mode of the fragment, which is vital for subsequent rational compound evolution. The routine use of the method depends on the availability of suitable fragment libraries, dedicated means to handle large numbers of samples, state-of-the-art synchrotron beamlines for fast diffraction measurements and largely automated solutions for the analysis of the results.
Here, the complete practical workflow and the included tools on how to conduct crystallographic fragment screening (CFS) at the Helmholtz-Zentrum Berlin (HZB) are presented. Preceding this workflow, crystal soaking conditions as well as data collection strategies are optimized for reproducible crystallographic experiments. Then, typically in a one to two-day procedure, a 96-membered CFS-focused library provided as dried ready-to-use plates is employed to soak 192 crystals, which are then flash-cooled individually. The final diffraction experiments can be performed within one day at the robot-mounting supported beamlines BL14.1 and BL14.2 at the BESSY&#160; II electron storage ring operated by the HZB in Berlin-Adlershof (Germany). Processing of the crystallographic data, refinement of the protein structures, and hit identification is fast and largely automated using specialized software pipelines on dedicated servers, requiring little user input.
Using the CFS workflow at the HZB enables routine screening experiments. It increases the chances for successful identification of fragment hits as starting points to develop more potent binders, useful for pharmacological or biochemical applications.

Crystallographic fragment screening at the Helmholtz-Zentrum Berlin is performed using a workflow with dedicated compound libraries, crystal handling tools, fast data collection facilities&#160;and largely automated data analysis. The presented protocol intends to maximize the output of such experiments to provide promising starting points for downstream structure-based ligand design.

Fragment screening is a technique that helps to identify promising starting points for ligand design. Given that crystals of the target protein are ...

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 ...

Preparation of Sample Support Films in Transmission Electron Microscopy using a Support Floatation Block

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant bottleneck. Macromolecular samples are ideally imaged directly from random orientations in a thin layer of vitreous ice. However, many samples are refractory to this, and protein denaturation at the air-water interface is a common problem. To overcome such issues, support films-including amorphous carbon, graphene, and graphene oxide-can be applied to the grid to provide a surface which samples can populate, reducing the probability of particles experiencing the deleterious effects of the air-water interface. The application of these delicate supports to grids, however, requires careful handling to prevent breakage, airborne contamination, or extensive washing and cleaning steps. A recent report describes the development of an easy-to-use floatation block that facilitates wetted transfer of support films directly to the sample. Use of the block minimizes the number of manual handling steps required, preserving the physical integrity of the support film, and the time over which hydrophobic contamination can accrue, ensuring that a thin film of ice can still be generated. This paper provides step-by-step protocols for the preparation of carbon, graphene, and graphene oxide supports for EM studies.

Sample preparation for cryo-electron microscopy (cryo-EM) is a significant bottleneck in the structure determination workflow of this method. Here, we provide detailed methods for using an easy-to-use, three-dimensionally printed block for the preparation of support films to stabilize samples for transmission EM studies.

Structure determination by cryo-electron microscopy (cryo-EM) has rapidly grown in the last decade; however, sample preparation remains a significant ...

Large-Scale Multi-Omics Genome-Wide Association Studies (Mo-GWAS): Guidelines for Sample Preparation and Normalization

Both gas chromatography-mass spectrometry (GC-MS) and liquid chromatography-mass spectrometry (LC-MS) are widely used metabolomics approaches to detect and quantify hundreds of thousands of metabolite features. However, the application of these techniques to a large number of samples is subject to more complex interactions, particularly for genome-wide association studies (GWAS). This protocol describes an optimized metabolic workflow, which combines an efficient and fast sample preparation with the analysis of a large number of samples for legume crop species. This slightly modified extraction method was initially developed for the analysis of plant and animal tissues and is based on extraction in methyl tert-butyl ether: methanol solvent to allow the capture of polar and lipid metabolites. In addition, we provide a step-by-step guide for reducing analytical variations, which are essential for the high-throughput evaluation of metabolic variance in GWAS.

Large-Scale Multi-Omics Genome-Wide Association Studies Mo-GWAS: Guidelines for Sample Preparation and Normalization

In this protocol, we present an optimized workflow, which combines an efficient and fast sample preparation of many samples. In addition, we provide a step-by-step guide to reduce analytical variations for high-throughput evaluation of metabolic GWAS studies.

Both gas chromatography-mass spectrometry (GC-MS) and liquid chromatography-mass spectrometry (LC-MS) are widely used metabolomics approaches to detect ...

Sample Preparation using a Lipid Monolayer Method for Electron Crystallographic Studies

Summary

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