The Gonen lab is supported by funds from the Howard Hughes Medical Institute. This study was supported by the National Institutes of Health P41GM136508.

1. Preparing small molecule samples
<ol>
	<li>Transfer a small amount (0.01 - 1 mg) of powder, liquid, or solids into a small vial or tube.</li>
	<li>For samples already in powder form, seal the tube using the cap until the sample is needed. Dry the liquid samples into powders prior to attempts at method 1 (step 3) or 2 (step 4). 
	&#8203;NOTE: Samples dissolved in liquid may use method 3 (5.X) below</li>
</ol>
2. Preparing TEM grids
NOTE: Some TEMs with autoloader systems require that the grids be clipped and placed into a cassette prior to loading into the TEM column. Clipping involves physically securing the 3 mm TEM grid into a metal ring that the autoloader can manipulate. This step and subsequent steps can be performed using either normal TEM grids, or TEM grids that have been clipped. For these experiments, it is often easier to manipulate the grids if they have been clipped ahead of time.
<ol>
	<li>Wrap plastic film around one end of a glass cover slide.</li>
	<li>Place the TEM grids onto the film on the top of the cover slide with the carbon side facing up. Identify the two sides of the grid under a light. The copper side shines and appears metallic, whereas the carbon side appears a drab, brown color (Figure 1C,D). For clipped grids, the carbon side should face the flat face of the clip ring.</li>
	<li>Place the slide with grids into the glow discharge chamber. Glow discharge the coverside for approximately 30 s using the negative setting at 15 pA. Store the grids on the cover slide inside of a glass Petri dish lined with filter paper prior to adding sample to the grids.</li>
</ol>
3. Applying sample to grids by creating a homogenous fine powder (Method 1)
<ol>
	<li>Remove a glow discharged TEM grid from the covered Petri dish using tweezers. Place the grid onto a circular filter paper with the carbon side facing up.</li>
	<li>Using a small spatula, remove a very small scoop of powder (approximately 0.1 mg) and place it onto a small, square, glass coverslip just next to the TEM grid on the filter paper. Place another small square glass slide or coverslip on top of the powder.</li>
	<li>With fingers, gently rub the two glass slides together to make a fine powder.</li>
	<li>Angle the coverslips and position them just above the TEM grid on the filter paper and continue to rub the coverslips together, just a few cm above the glow discharged TEM grid (Figure 1).</li>
	<li>Observe to see if the powder is falling towards the grid. Uncover the finely ground powder by removing one of the two glass coverslips. Gently brush the fine powder off of the coverslip using a piece of filter paper onto the TEM grid (Figure 1).</li>
</ol>
4. Applying sample to grids by applying the &#34;shaking&#34; method (Method 2)
<ol>
	<li>Grab a grid using a pair of tweezers and drop it into the vial or tube of powder sample. Close the vial using the plastic cap to ensure no material will escape when shaken.</li>
	<li>Grabbing the vial in the hand, shake the vial such that the powder and grid are both moving for approximately 10 - 30 s.</li>
	<li>Empty the vial contents onto a circular filter paper. Grabbing the grid on an edge, gently tap the grid edge on the filter paper to remove any excess material from the grid. 
	&#8203;NOTE: Depending on the type of vial and size, one may also use tweezers to remove the TEM grid without emptying the contents.</li>
</ol>
5. Applying sample to grids using the evaporation method (Method 3)
<ol>
	<li>Place a grid (clipped or not) onto the center of a circular piece of filter paper with the carbon side facing up and the copper side facing down.</li>
	<li>Using a 10 &#181;L gas-tight syringe, apply a small drop (approximately 1 - 3 &#181;L) of dissolved compound onto the carbon side of the grid.</li>
	<li>Gently move the filter paper with grids into a vacuum desiccation chamber. Cover the chamber and turn on the vacuum. Leave the grids under vacuum to dry for up to one day.</li>
	<li>Turn off the vacuum and allow the chamber to vent for 5 min. Venting the chamber avoids the grids moving or flying away when it is uncovered.</li>
</ol>
6. Freezing and loading grids into the TEM
<ol>
	<li>Grab the edge of the TEM grid using a set of tweezers, assuring that the tips do not puncture any of the grid squares. Lift the grid 1 - 2 cm above the filter paper and angle the grid at 90&#176; to the paper below. Gently tap the tweezers while keeping the grid firmly tweezed to remove any loose powder.</li>
	<li>Freeze the grid by moving the tip of the tweezers with the grid directly into a liquid nitrogen container by hand. This container is typically the grid loading station for the TEM, but can be any safe container, such as a liquid nitrogen safe thermos, is acceptable for transferring or storage. Liquid nitrogen is -196 &#176;C.</li>
	<li>Wait until the grid and tweezers stop boiling before further manipulations.</li>
	<li>Under liquid nitrogen or in nitrogen vapors, place the grid in the sample holder with the carbon side oriented such that the sample will be hit by the beam prior to the carbon support film.</li>
	<li>Load the sample holder into the TEM assuring that the grid is kept at liquid nitrogen temperatures at all times.
	<ol>
		<li>For autoloader systems, place the clipped grids into a cassette in a liquid nitrogen cooled container. This cassette transferred into a shuttling container than allows the autoloader robotics to accept the cassette while keeping the samples safe for shuttling between the autoloader and the column.</li>
		<li>For side-entry TEM setups, secure the grid to a commercial side-entry TEM holder. These holders have a sample preparation container that is filled with liquid nitrogen to allow transfer of the grid to the holder without warming the sample. The side-entry holder is inserted into the TEM directly with the sample secured at the end.</li>
	</ol>
	</li>
</ol>
7. Collecting MicroED data
<ol>
	<li>Locating and screening nanocrystals
	<ol>
		<li>Open the TEM column valves. Adjust the magnification using the hand panels to the lowest magnification possible. Find the beam by adjusting the intensity knob on the hand panels such that a round, bright area is visible on the fluorescent screen.</li>
		<li>Take an all-grid atlas at a low magnification (50 - 300x) using appropriate software14,15,16&#160;(Figure 2). Ensure that the microscope is well aligned for both low and high magnification imaging prior to collecting high-resolution MicroED data.</li>
		<li>Identify grid squares without broken carbon and have a visible black or dark material/grains on the film (Figure 2). Navigate around the grid, either physically using the joystick on the hand panels, or virtually on the collected Atlas, in order to search for grid squares that are not broken and contain microcrystals.</li>
		<li>Add the center of each of these squares to a list of grid locations for investigation. These locations can be added to a notebook, in the microscope user interface, or in microscope automation software.</li>
		<li>Increase the magnification to 500-1,300x and adjust the eucentric height at each stored grid location and update the saved Z value to the positions noted in 7.1.4.</li>
		<li>Search either on the fluorescent screen or on a fast camera at this higher magnification for small black spots/grains on the grid. A good sample with often have sharp edges at high magnification, suggesting crystalline order.</li>
		<li>Move a located potential crystal to the center of the screen and increase the magnification such that the TEM enters high magnification mode. 
		NOTE: This is referred to as either high-magnification, Zoom2, or SA mode on different instruments and typically corresponds to magnifications of 3,000x or higher.</li>
		<li>Insert a selected area aperture. Change the aperture to a larger or smaller size to assure that the selected area is just larger than the crystal (Figure 3).</li>
		<li>Switch to diffraction mode by pressing the diffraction button on the TEM hand panels, assuring that the fluorescent screen is inserted. Adjust the camera length using the magnification knob such that the edge is at least 1 &#197; resolution. 
		NOTE: Calibration of the diffraction lengths should be performed by a service engineer using a gold waffle grating or known specimen prior to attempting MicroED experiments.</li>
		<li>Adjust the diffraction focus such that the central spot is as sharp and small as possible. Using the diffraction shift knobs, move the central beam to the center of the fluorescent screen</li>
		<li>Insert the beam stop and make sure the beam is behind it. Lift the fluorescent screen. Take a short (approximately 1 s) exposure on the camera (Figure 4).</li>
		<li>Inspect the corresponding diffraction pattern. A good candidate for collecting a full dataset will have sharp spots that are regularly arranged in columns and rows, and the diffraction will extent to beyond 2 &#197;, preferably at least to 1 &#197; (Figure 4). Save the crystal coordinates in either the TEM user interface or by writing them down.</li>
		<li>Repeat diffraction screening for all the potential crystals of interest on the current grid square.</li>
	</ol>
	</li>
	<li>Data collection
	<ol>
		<li>Center a screened crystal on the screen at a high&#160;magnification (&#62; 1,000x). Adjust the eucentric height of the crystal using either an automatic routine or by hand.</li>
		<li>Insert the selected area aperture that best fits the crystal size and shape as determined in 7.1.8 (Figure 3). Tilt the stage in the negative and positive directions until the image is occluded by the grid bars (Figure 3). Note these angles for data collection purposes.</li>
		<li>Determine the number of frames that will be required to span the total angular wedge of the dataset. It is typical to use 0.5 - 1.0&#176; wedges for each frame, with frames being read out every 1 to 5s depending on the sensitivity of the camera. For example, a wedge spanning from -30&#176; to +30&#176;, rotating at a constant rate of 1&#176; s-1 with a frame read out every 1s, would require 60 total frames.</li>
		<li>Considering the maximum total tilt range of -72&#176; to +72&#176;, begin rotating the stage at a constant rate of 1&#176; s-1 and then begin reading the data out every 1s on a modern camera with a rolling shutter readout mode (Movie 1). This process can be performed manually or by using TEM specific software.
		<ol>
			<li>Set the rotation rate by specifying the % of the maximum tilt speed and when to stop in the microscope user interface or dedicated software. In this investigation, this was measured to be 0.3&#176; s-1 for each % of maximum speed but will need to be independently verified for each microscope. 
			NOTE: The tilting and camera data collection are set up independently here, but software programs14 can coordinate this to simplify the process.</li>
			<li>Discard approximately 1&#176; on each side of the specified wedge in most cases as dead time, where the stage is either ramping up or slowing down to the desired rotation speed.</li>
		</ol>
		</li>
		<li>Save data in a variety of formats as either individual frames or as a stack of images (Movie 1).</li>
		<li>Convert these diffraction frames to a typical crystallographic format using MicroED tools -available here (https://cryoem.ucla.edu). Diffraction images saved in SMV format are readily processed using documented&#160;crystallographic software17,18.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Shi, D., Nannenga, B. L., Iadanza, M. G., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+electron+crystallography+of+protein+microcrystals.">Three-dimensional electron crystallography of protein microcrystals.</a> eLife. 2, 01345 (2013).</li><li>Nannenga, B. L., Shi, D., Leslie, A. G. W., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+structure+determination+by+continuous-rotation+data+collection+in+MicroED.">High-resolution structure determination by continuous-rotation data collection in MicroED.</a> Nature Methods. 11 (9), 927-930 (2014).</li><li>Hattne, J., Martynowycz, M. W., Penczek, P. A., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroED+with+the+Falcon+III+direct+electron+detector.">MicroED with the Falcon III direct electron detector.</a> IUCrJ. 6 (5), 921-926 (2019).</li><li>Hattne, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroED+data+collection+and+processing.">MicroED data collection and processing.</a> Acta Crystallographica Section A Foundations and Advances. 71 (4), 353-360 (2015).</li><li>Henderson, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+potential+and+limitations+of+neutrons,+electrons+and+X-rays+for+atomic+resolution+microscopy+of+unstained+biological+molecules.">The potential and limitations of neutrons, electrons and X-rays for atomic resolution microscopy of unstained biological molecules.</a> Quarterly Reviews of Biophysics. 28 (2), 171-193 (1995).</li><li>Martynowycz, M. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroED+structure+of+the+human+adenosine+receptor+determined+from+a+single+nanocrystal+in+LCP.">MicroED structure of the human adenosine receptor determined from a single nanocrystal in LCP.</a> BioRxiv. , 316109 (2020).</li><li>Martynowycz, M. W., Zhao, W., Hattne, J., Jensen, G. J., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collection+of+continuous+rotation+MicroED+data+from+ion+beam-milled+crystals+of+any+size.">Collection of continuous rotation MicroED data from ion beam-milled crystals of any size.</a> Structure. 27 (3), 545-548 (2019).</li><li>Martynowycz, M. W., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ligand+incorporation+into+protein+microcrystals+for+MicroED+by+on-grid+soaking.">Ligand incorporation into protein microcrystals for MicroED by on-grid soaking.</a> Structure. , (2020).</li><li>Martynowycz, M. W., Khan, F., Hattne, J., Abramson, J., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroED+structure+of+lipid-embedded+mammalian+mitochondrial+voltage-dependent+anion+channel.">MicroED structure of lipid-embedded mammalian mitochondrial voltage-dependent anion channel.</a> Proceedings of the National Academy of Sciences. 117 (51), 32380-32385 (2020).</li><li>Jones, C. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+CryoEM+method+MicroED+as+a+powerful+tool+for+small+molecule+structure+determination.">The CryoEM method MicroED as a powerful tool for small molecule structure determination.</a> ACS Central Science. 4 (11), 1587-1592 (2018).</li><li>Dick, M., Sarai, N. S., Martynowycz, M. W., Gonen, T., Arnold, F. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tailoring+tryptophan+synthase+TrpB+for+selective+quaternary+carbon+bond+formation.">Tailoring tryptophan synthase TrpB for selective quaternary carbon bond formation.</a> Journal of the American Chemical Society. 141 (50), 19817-19822 (2019).</li><li>Gallagher-Jones, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sub-&#229;ngstr&#246;m+cryo-EM+structure+of+a+prion+protofibril+reveals+a+polar+clasp.">Sub-&#229;ngstr&#246;m cryo-EM structure of a prion protofibril reveals a polar clasp.</a> Nature Structural &amp; Molecular Biology. 25 (2), 131-134 (2018).</li><li>Ting, C. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+a+scaffold+peptide+in+the+biosynthesis+of+amino+acid-derived+natural+products.">Use of a scaffold peptide in the biosynthesis of amino acid-derived natural products.</a> Science. 365 (6450), 280-284 (2019).</li><li>de la Cruz, M. J., Martynowycz, M. W., Hattne, J., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroED+data+collection+with+SerialEM.">MicroED data collection with SerialEM.</a> Ultramicroscopy. 201, 77-80 (2019).</li><li>Mastronarde, D. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+electron+microscope+tomography+using+robust+prediction+of+specimen+movements.">Automated electron microscope tomography using robust prediction of specimen movements.</a> Journal of Structural Biology. 152 (1), 36-51 (2005).</li><li>Schorb, M., Haberbosch, I., Hagen, W. J. H., Schwab, Y., Mastronarde, D. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Software+tools+for+automated+transmission+electron+microscopy.">Software tools for automated transmission electron microscopy.</a> Nature Methods. 16 (6), 471-477 (2019).</li><li>Kabsch, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=XDS.">XDS.</a> Acta Crystallographica Section D Biological Crystallography. 66 (2), 125-132 (2010).</li><li>Winter, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DIALS:+Implementation+and+evaluation+of+a+new+integration+package.">DIALS: Implementation and evaluation of a new integration package.</a> Acta Crystallographica Section D. 74 (2), 85-97 (2018).</li><li>de la Cruz, M. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic-resolution+structures+from+fragmented+protein+crystals+with+the+cryoEM+method+MicroED.">Atomic-resolution structures from fragmented protein crystals with the cryoEM method MicroED.</a> Nature Methods. 14 (4), 399-402 (2017).</li><li>Shi, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+collection+of+MicroED+data+for+macromolecular+crystallography.">The collection of MicroED data for macromolecular crystallography.</a> Nature Protocols. 11 (5), 895-904 (2016).</li><li>Nannenga, B. L., Shi, D., Hattne, J., Reyes, F. E., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structure+of+catalase+determined+by+MicroED.">Structure of catalase determined by MicroED.</a> eLife. 3, 03600 (2014).</li><li>Martynowycz, M. W., Zhao, W., Hattne, J., Jensen, G. J., Gonen, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Qualitative+Analyses+of+Polishing+and+Precoating+FIB+Milled+Crystals+for+MicroED.">Qualitative Analyses of Polishing and Precoating FIB Milled Crystals for MicroED.</a> Structure. 27 (10), 1594-1600 (2019).</li></ol>

The authors have nothing to disclose.

Sample preparation is typically an iterative process, where optimizations are made after sessions of screening and data collection. For small-molecule samples, it is often prudent to first attempt grid preparation without glow-discharging the grids, since many pharmaceuticals tend to be hydrophobic10,11. If the grids have too few nanocrystalline deposits, it is a good idea to try again after first glow-discharging the grids. It may be the case that the crystals f...

Microcrystal electron diffraction (MicroED) is an electron cryo-microscopy (cryo-EM) method for determining atomic resolution structures from sub-micrometer sized crystals1,2. Crystals are applied to standard transmission electron microscope (TEM) grids and frozen by either plunging into liquid ethane or liquid nitrogen. Grids are then loaded into a TEM operating at cryogenic temperatures. Crystals are located on the grid and screened for initial diffraction quality. Continuous rotation MicroED data are collected from a subset of the screened crystals, where the data are saved using a fast camera as a movie3. These movies are converted to a standard crystallographic format and processed almost identically as an X-ray crystallography experiment4.
MicroED was originally developed to investigate protein microcrystals1,2. A bottleneck in protein crystallography is growing large, well-ordered crystals for traditional synchrotron X-ray diffraction experiments. As electrons interact with matter orders of magnitude stronger than X-rays, the limitations of the crystal size needed to produce detectable diffraction is considerably smaller5. Additionally, the ratio of elastic to inelastic scattering events is more favorable for electrons, suggesting that more useful data can be collected with a smaller overall exposure5. Constant developments have allowed for MicroED data to be collected from the most challenging microcrystals6,7,8,9.
Recently, MicroED has been shown to be a powerful tool for determining the structures of small molecule pharmaceuticals from apparently amorphous materials10,11,12,13. These powders can come straight from a bottle of purchased reagent, a purification column, or even from crushing a pill into a fine powder10. These powders appear amorphous by eye, but may be either entirely composed of nanocrystals or merely contain trace amounts of nanocrystalline deposits in a greater non-crystalline, amorphous fraction. Application of the material to the grid is facile, and the subsequent steps of crystal identification, screening, and data collection might even be automated in the near future14. While others may use different methods for sample preparation and data collection, here the protocols developed and used in the Gonen laboratory for preparing samples of small molecules for MicroED and for data collection are detailed.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.1-1.5mL Eppendorf tubes</td>
			<td>Fisher Scientific</td>
			<td>14-282-300</td>
			<td>Any vial or tube will do.</td>
		</tr>
		<tr>
			<td>Autogrid clips</td>
			<td>Thermo-Fisher</td>
			<td>1036173</td>
			<td>Clipped grids are not required for MicroED. They are required for Thermo-Fisher TEMs equipped with an autoloader system.</td>
		</tr>
		<tr>
			<td>Autogrid C-rings</td>
			<td>Thermo-Fisher</td>
			<td>1036171</td>
			<td></td>
		</tr>
		<tr>
			<td>Carbamazapine</td>
			<td>Sigma</td>
			<td>C4024-1G</td>
			<td>Any amount will suffice for these experiments</td>
		</tr>
		<tr>
			<td>CMOS based detector</td>
			<td>Thermo-Fisher</td>
			<td>CetaD 16M</td>
			<td>We used a CetaD 16M, but any detector with rolling shutter mode or sufficiently fast readout is acceptable.&#160;</td>
		</tr>
		<tr>
			<td>Delphi software</td>
			<td>Thermo-Fisher</td>
			<td>N/A</td>
			<td>Software on Thermo-Fisher TEM systems that allows for manual rotation of the sample stage</td>
		</tr>
		<tr>
			<td>EPU-D software</td>
			<td>Thermo-Fisher</td>
			<td>N/A</td>
			<td>Commercial software for the acquisition of MicroED data</td>
		</tr>
		<tr>
			<td>Glass cover slides</td>
			<td>Hampton</td>
			<td>HR3-231</td>
			<td></td>
		</tr>
		<tr>
			<td>Glow discharger</td>
			<td>Pelco</td>
			<td>easiGlow</td>
			<td></td>
		</tr>
		<tr>
			<td>High PrecisionTweezers</td>
			<td>EMS</td>
			<td>78325-AC</td>
			<td>Any high precision tweezer will do</td>
		</tr>
		<tr>
			<td>Liquid nitrogen vessel</td>
			<td>Spear Lab</td>
			<td>FD-800</td>
			<td>A standard foam vessel for handling specimens under liquid nitrogen - 800mL</td>
		</tr>
		<tr>
			<td>SerialEM software</td>
			<td>UC Boulder</td>
			<td>N/A</td>
			<td>Free software distributed by D. Mastronarde. Department of Molecular, Cellular, and Developmental Biology</td>
		</tr>
		<tr>
			<td>TEM grids</td>
			<td>Quantifoil/EMS</td>
			<td>Q310CMA</td>
			<td>Multi-A 300 mesh grids were used here, but any thin carbon grids will work. For these small molecules, we suggest starting with continuous carbon.&#160;</td>
		</tr>
		<tr>
			<td>transmission electron microscope (TEM)</td>
			<td>Thermo-Fisher</td>
			<td>Talos Arctica</td>
			<td></td>
		</tr>
		<tr>
			<td>Whatman circular filter paper</td>
			<td>Millipore-Sigma</td>
			<td>WHA1001090</td>
			<td>90mm or larger</td>
		</tr>
	</tbody>
</table>

microcrystal electron diffraction of small molecules

A detailed protocol for preparing small molecule samples for microcrystal electron diffraction (MicroED) experiments is described. MicroED has been developed to solve structures of proteins and small molecules using standard electron cryo-microscopy (cryo-EM) equipment. In this way, small molecules, peptides, soluble proteins, and membrane proteins have recently been determined to high resolutions. Protocols are presented here for preparing grids of small-molecule pharmaceuticals using the drug carbamazepine as an example. Protocols for screening and collecting data are presented. Additional steps in the overall process, such as data integration, structure determination, and refinement are presented elsewhere. The time required to prepare the small-molecule grids is estimated to be less than 30 min.

MicroED is a cryoEM method that leverages the strong interactions between electrons and matter, which allows for the investigation of vanishingly small crystals12,13. After these steps, it is expected to have a diffraction movie in crystallographic format collected from microcrystals (Movie 1). Here, the technique is demonstrated using carbamazepine12. The results show a continuous rotation MicroED dataset from a carbamaze...

Watch this Scientific Journal Video about Microcrystal Electron Diffraction of Small Molecules at JoVE.com

Microcrystal Electron Diffraction of Small Molecules

Here, we describe the procedures developed in our laboratory for preparing powders of small molecule crystals for microcrystal electron diffraction (MicroED) experiments.

A detailed protocol for preparing small molecule samples for microcrystal electron diffraction (MicroED) experiments is described. MicroED has been ...

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6.5K Views.  University of California Los Angeles. Here, we describe the procedures developed in our laboratory for preparing powders of small molecule crystals for microcrystal electron diffraction (MicroED) experiments. 

Video: Microcrystal Electron Diffraction of Small Molecules

Microcrystallography of Protein Crystals and In Cellulo Diffraction

The advent of high-quality microfocus beamlines at many synchrotron facilities has permitted the routine analysis of crystals smaller than 10 &#181;m in their largest dimension, which used to represent a challenge. We present two alternative workflows for the structure determination of protein microcrystals by X-ray crystallography with a particular focus on crystals grown in vivo. The microcrystals are either extracted from cells by sonication and purified by differential centrifugation, or analyzed in cellulo after cell sorting by flow cytometry of crystal-containing cells. Optionally, purified crystals or crystal-containing cells are soaked in heavy atom solutions for experimental phasing. These samples are then prepared for diffraction experiments in a similar way by application onto a micromesh support and flash cooling in liquid nitrogen. We briefly describe and compare serial diffraction experiments of isolated microcrystals and crystal-containing cells using a microfocus synchrotron beamline to produce datasets suitable for phasing, model building and refinement.
These workflows are exemplified with crystals of the Bombyx mori cypovirus 1 (BmCPV1) polyhedrin produced by infection of insect cells with a recombinant baculovirus. In this case study, in cellulo analysis is more efficient than analysis of purified crystals and yields a structure in ~8 days from expression to refinement.

Microcrystallography of Protein Crystals and In Cellulo Diffraction

A protocol is presented for X-ray crystallography using protein microcrystals. Two examples analyzing in vivo-grown microcrystals after purification or in cellulo are compared.

The advent of high-quality microfocus beamlines at many synchrotron facilities has permitted the routine analysis of crystals smaller than 10 &#181;m in ...

Force Spectroscopy of Single Protein Molecules Using an Atomic Force Microscope

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. Atomic force microscopy (AFM) can address this problem by enabling stretching and relaxation of single protein molecules, which gives direct evidence of specific unfolding and refolding characteristics. AFM-based single-molecule force-spectroscopy (AFM-SMFS) provides a means to consistently measure high-energy conformations in proteins that are not possible in traditional bulk (biochemical) measurements. Although numerous papers were published to show principles of AFM-SMFS, it is not easy to conduct SMFS experiments due to a lack of an exhaustively complete protocol. In this study, we briefly illustrate the principles of AFM and extensively detail the protocols, procedures, and data analysis as a guideline to achieve good results from SMFS experiments. We demonstrate representative SMFS results of single protein mechanical unfolding measurements and we provide troubleshooting strategies for some commonly encountered problems.

We describe the detailed procedures and strategies to measure the mechanical properties and mechanical unfolding pathways of single protein molecules using an atomic force microscope. We also show representative results as a reference for selection and justification of good single protein molecule recordings.

The determination of the folding process of proteins from their amino acid sequence to their native 3D structure is an important problem in biology. ...

Semi-automated Biopanning of Bacterial Display Libraries for Peptide Affinity Reagent Discovery and Analysis of Resulting Isolates

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. Peptide affinity reagents can be used for similar applications to antibodies, including sensing and therapeutics, but are more robust and able to perform in more extreme environments. Specific enrichment of peptide capture agents to a protein target of interest is enhanced using semi-automated sorting methods which improve binding and wash steps and therefore decrease the occurrence of false positive binders. A semi-automated sorting method is described herein for use with a commercial automated magnetic-activated cell sorting device with an unconstrained bacterial display sorting library expressing random 15-mer peptides. With slight modifications, these methods are extendable to other automated devices, other sorting libraries, and other organisms. A primary goal of this work is to provide a comprehensive methodology and expound the thought process applied in analyzing and minimizing the resulting pool of candidates. These techniques include analysis of on-cell binding using fluorescence-activated cell sorting (FACS), to assess affinity and specificity during sorting and in comparing individual candidates, and the analysis of peptide sequences to identify trends and consensus sequences for understanding and potentially improving the affinity to and specificity for the target of interest.

Biopanning bacterial display libraries is a proven technique for discovery of peptide affinity reagents, a robust alternative to antibodies. The semi-automated sorting method herein has streamlined biopanning to decrease the occurrence of false positives. Here we illustrate the thought process and techniques applied in evaluating candidates and minimizing downstream analysis.

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. ...

Capturing the Interaction Kinetics of an Ion Channel Protein with Small Molecules by the Bio-layer Interferometry Assay

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe the application of this novel label-free technique to study the interaction of human EAG1 (hEAG1) channel proteins with the small molecule PIP2. hEAG1 channel has been recognized as potential therapeutic target because of its aberrant overexpression in cancers and a few gain-of-function mutations involved in some types of neurological diseases. We purified&#160;hEAG1 channel proteins from a mammalian stable expression system and measured the interaction with PIP2 by BLI. The successful measurement of the kinetics of binding between hEAG1 protein and PIP2 demonstrates that the BLI assay is a potential high-throughput approach used for novel small-molecule ligand screening in ion channel pharmacology.

The protocol here describes the interactions of purified hEAG1 ion channel protein with the small molecule lipid ligand phosphatidylinositol 4, 5-bisphosphate (PIP2). The measurement demonstrates that BLI could be a potential method for novel small-molecule ion channel ligand screening.

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe ...

Analysis of &#946;-Amyloid-induced Abnormalities on Fibrin Clot Structure by Spectroscopy and Scanning Electron Microscopy

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and structure by spectrometry and scanning electron microscopy (SEM). A&#946;, which forms neurotoxic amyloid aggregates in Alzheimer&#39;s disease (AD), has been shown to interact with fibrinogen. This A&#946;-fibrinogen interaction makes the fibrin clot structurally abnormal and resistant to fibrinolysis. A&#946;-induced abnormalities in fibrin clotting may also contribute to cerebrovascular aspects of the AD pathology such as microinfarcts, inflammation, as well as, cerebral amyloid angiopathy (CAA). Given the potentially critical role of neurovascular deficits in AD pathology, developing compounds which can inhibit or lessen the A&#946;-fibrinogen interaction has promising therapeutic value. In vitro methods by which fibrin clot formation can be easily and systematically assessed are potentially useful tools for developing therapeutic compounds. Presented here is an optimized protocol for in vitro generation of the fibrin clot, as well as analysis of the effect of A&#946; and A&#946;-fibrinogen interaction inhibitors. The clot turbidity assay is rapid, highly reproducible and can be used to test multiple conditions simultaneously, allowing for the screening of large numbers of A&#946;-fibrinogen inhibitors. Hit compounds from this screening can be further evaluated for their ability to ameliorate A&#946;-induced structural abnormalities of the fibrin clot architecture using SEM. The effectiveness of these optimized protocols is demonstrated here using TDI-2760, a recently identified A&#946;-fibrinogen interaction inhibitor.

Presented here are two methods that can be used individually or in combination to analyze the effect of beta-amyloid on fibrin clot structure. Included is a protocol for creating an in vitro fibrin clot, followed by clot turbidity and scanning electron microscopy methods.

This article presents methods for generating in vitro fibrin clots and analyzing the effect of beta-amyloid (A&#946;) protein on clot formation and ...

Single-Molecule Tracking Microscopy - A Tool for Determining the Diffusive States of Cytosolic Molecules

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and millisecond temporal resolution. These capabilities make single-molecule localization microscopy ideally suited to study molecular level biological functions in physiologically relevant environments. Here, we demonstrate an integrated protocol for both acquisition and processing/analysis of single-molecule tracking data to extract the different diffusive states a protein of interest may exhibit. This information can be used to quantify molecular complex formation in living cells. We provide a detailed description of a camera-based 3D single-molecule localization experiment, as well as the subsequent data processing steps that yield the trajectories of individual molecules. These trajectories are then analyzed using a numerical analysis framework to extract the prevalent diffusive states of the fluorescently labeled molecules and the relative abundance of these states. The analysis framework is based on stochastic simulations of intracellular Brownian diffusion trajectories that are spatially confined by an arbitrary cell geometry. Based on the simulated trajectories, raw single-molecule images are generated and analyzed in the same way as experimental images. In this way, experimental precision and accuracy limitations, which are difficult to calibrate experimentally, are explicitly incorporated into the analysis workflow. The diffusion coefficient and relative population fractions of the prevalent diffusive states are determined by fitting the distributions of experimental values using linear combinations of simulated distributions. We demonstrate the utility of our protocol by resolving the diffusive states of a protein that exhibits different diffusive states upon forming homo- and hetero-oligomeric complexes in the cytosol of a bacterial pathogen.

3D single-molecule localization microscopy is utilized to probe the spatial positions and motion trajectories of fluorescently labeled proteins in living bacterial cells. The experimental and data analysis protocol described herein determines the prevalent diffusive behaviors of cytosolic proteins based on pooled single-molecule trajectories.

Single-molecule localization microscopy probes the position and motions of individual molecules in living cells with tens of nanometer spatial and ...

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Micropatterning Transmission Electron Microscopy Grids to Direct Cell Positioning within Whole-Cell Cryo-Electron Tomography Workflows

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules present in the cellular context and preserved in a near-native frozen-hydrated state. However, there are challenges associated with culturing and/or adhering cells onto TEM grids in a manner that is suitable for tomography while retaining the cells in their physiological state. Here, a detailed step-by-step protocol is presented on the use of micropatterning to direct and promote eukaryotic cell growth on TEM grids. During micropatterning, cell growth is directed by depositing extra-cellular matrix (ECM) proteins within specified patterns and positions on the foil of the TEM grid while the other areas remain coated with an anti-fouling layer. Flexibility in the choice of surface coating and pattern design makes micropatterning broadly applicable for a wide range of cell types. Micropatterning is useful for studies of structures within individual cells as well as more complex experimental systems such as host-pathogen interactions or differentiated multi-cellular communities. Micropatterning may also be integrated into many downstream whole-cell cryo-ET workflows, including correlative light and electron microscopy (cryo-CLEM) and focused-ion beam milling (cryo-FIB).

The goal of this protocol is to direct cell adhesion and growth to targeted areas of grids for cryo-electron microscopy. This is achieved by applying an anti-fouling layer that is ablated in user-specified patterns followed by deposition of extra-cellular matrix proteins in the patterned areas prior to cell seeding.

Whole-cell cryo-electron tomography (cryo-ET) is a powerful technology that is used to produce nanometer-level resolution structures of macromolecules ...

On-Chip Crystallization and Large-Scale Serial Diffraction at Room Temperature

Biochemical reactions and biological processes can be best understood by demonstrating how proteins transition among their functional states. Since cryogenic temperatures are non-physiological and may prevent, deter, or even alter protein structural dynamics, a robust method for routine X-ray diffraction experiments at room temperature is highly desirable. The crystal-on-crystal device and its accompanying hardware and software used in this protocol are designed to enable in situ X-ray diffraction at room temperature for protein crystals of different sizes without any sample manipulation. Here we present the protocols for the key steps from device assembly, on-chip crystallization, optical scanning, crystal recognition to X-ray shot planning and automated data collection. Since this platform requires no crystal harvesting nor any other sample manipulation, hundreds to thousands of protein crystals grown on chip can be introduced into an X-ray beam in a programmable and high-throughput manner.

This contribution describes how to set up protein crystallization on crystal-on-crystal devices and how to perform automated serial data collection at room temperature using the on-chip crystallization platform.

Biochemical reactions and biological processes can be best understood by demonstrating how proteins transition among their functional states. Since ...

The Extraction of Liver Glycogen Molecules for Glycogen Structure Determination

Liver glycogen is a hyperbranched glucose polymer that is involved in the maintenance of blood sugar levels in animals. The properties of glycogen are influenced by its structure. Hence, a suitable extraction method that isolates representative samples of glycogen is crucial to the study of this macromolecule. Compared to other extraction methods, a method that employs a sucrose density gradient centrifugation step can minimize molecular damage. Based on this method, a recent publication describes how the density of the sucrose solution used during centrifugation was varied (30%, 50%, 72.5%) to find the most suitable concentration to extract glycogen particles of a wide variety of sizes, limiting the loss of smaller particles. A 10 min boiling step was introduced to test its ability to denature glycogen degrading enzymes, thus preserving glycogen. The lowest sucrose concentration (30%) and the addition of the boiling step were shown to extract the most representative samples of glycogen.

An optimal sucrose concentration was determined for the extraction of liver glycogen using sucrose density gradient centrifugation. The addition of a 10 min boiling step to inhibit glycogen-degrading enzymes proved beneficial.

Liver glycogen is a hyperbranched glucose polymer that is involved in the maintenance of blood sugar levels in animals. The properties of glycogen are ...