We thank the kind donations of AAV5 sample by Mavis Agbandje-McKenna and Antonette Bennett, and of TMV sample by Ruben Diaz. This work is supported by American Heart Association grant 14GRNT19660003 and VCU startup funds to MS.

1. Cryo-holder Preparation and Maintenance
NOTE: The dry pumping station, consisting of a turbo pumping station and one or more cold stage controllers, is primarily used for three purposes: a) It is a safe place to store the cryo holders when they are not being used. The proper way to store the holders is to leave them in the station under vacuum. b) To evaporate the frost and humidity buildup that occurs when the liquid nitrogen is removed from the dewar and the tip of the cryo holder is exposed; this is achieved with the warm-up cycle. c) To regenerate the zeolite desiccant, and to achieve a high vacuum in the cryoholder dewars. This is necessary for holding a constant temperature when doing cryo electron microscopy.
<ol>
	<li>Warm-up Cycle.
	<ol>
		<li>Insert the cryo holder into one of the pumping ports of the pumping station. Hook up one of the evacuation tubes to the metal outlet underneath the vacuum valve of the holder. Make sure that all valves on the station (V1, V2, and V3) and on the cryo holder are closed.</li>
		<li>Turn on the cold stage controller and connect it to the holder through the control cord. Turn on the power switch of the turbo pumping station.</li>
		<li>Start the Warm-up cycle option in the cold stage controller. When the vacuum in the turbo pumping station reads 10-3 Torr or better, open the butterfly valve V2, wait until the vacuum stabilizes, and then open the butterfly valve V1.</li>
		<li>Run the warm up cycle for around 30 min until the temperature in the holder is stable above 20 &#176;C. Close V1, and then stop the cycle.</li>
	</ol>
	</li>
	<li>Zeolite cycle. Run the Zeolite Cycle Between 4 hr and O/N Before a CryoEM Session, and Once a Week if Not in Use.
	<ol>
		<li>Start the Zeolite cycle option and select the necessary time to attain a good vacuum, in the range of 1-2 x 10-4&#160;Torr, and thus long hold time. When the vacuum reaches 10-3 Torr, open the dewar evacuation valve, V3. The pumping station will then evacuate the line to the holder; wait until the vacuum stabilizes.</li>
		<li>Open the valve on the cryo holder. When the cycle is complete, close the valves in the opposite order in which they were opened: the valve on the holder, then V3, V2, and finally V1. Turn off the turbo pumping station and the cold stage controller, and disconnect the cryo holder from the cold stage controller and the turbo pumping station.</li>
	</ol>
	</li>
</ol>
2. Grid Preparation
<ol>
	<li>Cleaning. 
	NOTE: When using commercial holey grids, it is recommended to clean them before use, in order to remove any residual polymer that was used in the fabrication process. Do this in a glass Petri dish with 5 layers of filter paper laid at the bottom.
	<ol>
		<li>Place the grids on the filter paper with the holey carbon side facing up.</li>
		<li>Using a glass Pasteur pipette, wet the paper with several drops of chloroform around the grids until they just start floating.</li>
		<li>Cover the Petri dish with the lid slightly open in a fume hood O/N.</li>
	</ol>
	</li>
	<li>NOTE: the thin carbon support (steps 2.2-2.3) can be omitted as the ice layer can be directly formed across the small holes in the holey carbon grid.
	<ol>
		<li>Using a pair of tweezers, cleave a piece of mica to expose two new surfaces. Place the new exposed surfaces of mica face-up on a white piece of paper in a Petri dish.</li>
		<li>Place the Petri dish in the bell jar of a carbon evaporator. Start the pump down sequence until vacuum is below 2 x 10-6 Torr. To evaporate any impurities on the surface of the carbon rod, perform a pre-evaporation with the Petri dish covered. Then air the bell jar and remove the cover from the Petri dish.</li>
		<li>Perform the pump down sequence until the vacuum is below 10-6 Torr and coat the mica with a thin layer (~5 nm) of carbon. Use eye protection during carbon evaporation. Monitor the thickness of the carbon film by the resulting darkness on the paper that had been placed under the mica sheets.</li>
	</ol>
	</li>
	<li>Transfer the Thin Carbon to the TEM Grid.
	<ol>
		<li>Cut a 0.5 x 1 cm piece of coated mica and float the carbon off onto the flat surface of filtered, de-ionized water. Use optical lens paper to get a flat water surface. Using the tweezers, put the holey carbon side of the grid in contact with the upper surface of the floating carbon layer and pick it up.</li>
		<li>Repeat Steps 2.3.1 and 2.3.2 until a sufficient number of grids are prepared.</li>
	</ol>
	</li>
	<li>Glow Discharge. 
	NOTE: The glow discharge converts the naturally hydrophobic carbon-coating layer of the grids into hydrophilic. This step is optional.
	<ol>
		<li>Place the grids with the carbon side facing up on a 7.5 x 2.5 cm glass slide covered with Parafilm. Place it into the chamber of the glow-discharge equipment and close the chamber.</li>
		<li>Pump the bell jar and glow-discharge for 20 sec at 25 mA and 7 x 10-2 mbars. 
		NOTE: During this time a light purple glow becomes visible. Remove the glass slide with the grids and use them within 1 hr, otherwise, they will need to be glow discharged again.</li>
	</ol>
	</li>
</ol>
3. Sample Plunge-freezing
NOTE: Use safety glasses and appropriate footwear when using this instrument to protect against liquid nitrogen and ethane splashes.
<ol>
	<li>Turn on the plunge-freezing apparatus. Fill humidifier reservoir with distilled water. Set the desired temperature, relative humidity and plunging conditions for blot time, blot force and adsorption time (22 &#176;C, 95%, 2 sec, 2 and 40 sec in the example presented here). Place new blotting filter paper.</li>
	<li>Cool the ethane container down by filling the outer reservoir with liquid nitrogen until stability is achieved (about 10 min). Once the liquid nitrogen stops boiling, place the tip of the tube coming from the ethane tank in the inner chamber and open the valve very slowly. Liquefy the ethane in the inner chamber and fill it to 1 mm from the top. Avoid freezing the ethane. CAUTION: ethane is extremely flammable and explosive.</li>
	<li>Make sure tweezers&#160;are aligned within the plunge freezing apparatus.</li>
	<li>Turn on humidity. Load a grid onto the tweezers, and load 3-5 &#181;l of sample onto the carbon surface (dull side) of the holey grid. Wait for the sample to adsorb to the grid and perform the blotting to achieve a thin liquid layer.</li>
	<li>Flash-freeze the grid by plunging it into the liquid ethane. Remove the tweezers-grid assembly from the liquid ethane holding the tweezers steadily, and transfer it to the outer chamber with liquid nitrogen. Transfer the grid to a small grid box immersed in the liquid nitrogen. Make sure to keep the vitrified sample in liquid nitrogen&#160;all the time during transfer to either the storage tank or to the cryo holder. 
	NOTE: Grid boxes can be stored in a large capacity (35-50 L) liquid nitrogen dewar for at least 1 year. For convenient storage they can be placed in a 50 ml Falcon tube with two drilled holes at the top and a fishing line attached to its cap that can be pulled from the mouth of the dewar.</li>
</ol>
4. Transfer the Cryo-grid to the TEM
<ol>
	<li>Insert the cryo holder into the cryo-transfer workstation. Take care not to damage the tip of the holder during insertion. Check for the presence of small fibers and dust on the rod and the O-ring of the cryo holder using a loupe, if there is any remove it with optical lens paper.</li>
	<li>Fill the dewar of the cryo holder and the workstation vessel with liquid nitrogen. Avoid spilling the liquid nitrogen by using the trapped funnel that comes with the workstation.</li>
	<li>Connect the cryo holder to the cold stage controller to monitor the temperature, and keep adding liquid nitrogen&#160;until the temperature stabilizes at around -194 &#176;C. Make sure that the level of liquid nitrogen is under the level of the vacuum seal both in the workstation vessel and the cryo holder dewar. Pre-cool the grid tweezers, the clip ring and the blunt end of the clip ring tool on the workstation. Also pre-cool a set of large tweezers and a screwdriver in another dewar filled with liquid nitrogen.</li>
	<li>Quickly transfer the cryo grid box containing the frozen hydrated grids to the liquid nitrogen in the workstation using the large tweezers. Open the cryo grid box with the screwdriver, take the frozen hydrated grid and place it into the sample holder slot.</li>
	<li>Place the clip ring on top of the grid and gently press it with the blunt end of the clip ring tool until it is firmly seated in place. Close the cryo-shutter. Remove the tools and cryo grid box from the workstation.</li>
	<li>Move the entire workstation with holder and grid to the microscope console in order to minimize the transfer time of the grid in room air.</li>
	<li>Top off the TEM anti-contaminator with liquid nitrogen, which had been previously filled and cooled down for at least 20 min. Pre-tilt the microscope stage to -60&#176;. 
	NOTE: This will minimize loss of liquid nitrogen as the holder is inserted into the airlock.</li>
	<li>Begin the pre-pump airlock cycle on the microscope. Make sure that the valve to the electron gun is closed (especially if it is an FEG TEM). Wait for the pre-pump airlock sequence to finish (about 2 min) before inserting the cryo holder. 
	NOTE: This is indicated by the red light on the stage being extinguished.</li>
	<li>Insert the holder into the airlock and rotate it 90&#176; clockwise;&#160;connect the cold stage controller cord to the cryo holder to monitor the temperature. Wait again until the red light is extinguished and rotate both the stage and&#160;the holder back into the 0&#176; position to insert the holder&#160;into the TEM&#8217;s high vacuum column.</li>
	<li>Make sure that the temperature never goes above -160 &#176;C to avoid formation of crystalline ice.</li>
	<li>Refill the dewar of the cryo holder. Wait 15 &#8211; 45 min until the holder has thermally equilibrated and the TEM vacuum has recovered before recording images.</li>
	<li>If bubbling of the liquid nitrogen is detected, change the liquid nitrogen&#160;filling height, or gently touch the bubbling origin with a cotton swab.</li>
</ol>
5. Low-dose Data Collection
NOTE: The low-dose data collection technique is used to minimize the electron radiation damage to the sample by limiting the exposure to 20 electrons/&#197;2. This is achieved by alternating between three configurations: &#8220;search&#8221;, with low magnification (~5,000X) and wide field of view, &#8220;focus&#8221;, with high magnification (~150,000X) and small field of view, and &#8220;exposure&#8221;, with the desired final magnification and containing the area of interest to be recorded. Blank the beam in between modes.

<ol>
	<li>Set Up the TEM and the Low-dose Program
	<ol>
		<li>Retract the cryo-shutter and open the column valves.</li>
		<li>Center the stage and set the eucentric height (Z) using the alpha wobbler to rock the stage &#177; 15&#176;. Adjust the Z position until there is no appreciable movement while the stage is rocking. Activate the low-dose program and select the detector for each mode.</li>
		<li>In exposure mode find an area of the grid that has no carbon, adjust the illumination by selecting the optimal condenser aperture and spot size so that the area to be recorded is fully illuminated. Be sure to spread the beam in the overcondensation direction.</li>
		<li>In search mode, find a small feature that will be easily identifiable at higher magnification. In exposure mode, center this feature with the joystick (the x-y stage controller). Start at lower magnification if the feature is not in the field of view.</li>
		<li>In search mode, bring this feature to the center of the field using the low dose x-y image shift controls. Go to exposure mode and move the feature along the tilt axis using the joystick until it is out of the area to be recorded (0.5 - 3 microns). Go to focus mode and center the feature using the x-y image shift controls. Start at lower magnification if the feature is not in the field of view.</li>
		<li>Keep the beam centered at all times.</li>
	</ol>
	</li>
	<li>Data Collection
	<ol>
		<li>Retract the cryo-shutter and open the column valves. Activate the low dose program.</li>
		<li>In search mode, identify a grid square containing thin, homogeneous ice.</li>
		<li>Select a suitable hole and place it in the center of the field. Switch to focus mode and focus the image. Then underfocus the image between 0.5 - 4.5 microns.</li>
		<li>Switch to exposure mode and record the image.</li>
		<li>Switch to search mode and repeat steps 5.2.3-5.2.4. Move across the grid square avoiding pre-exposing good holes to be recorded.</li>
		<li>Finish the grid square and move to another good one.</li>
		<li>Readjust height (Z) and continue data collection.</li>
	</ol>
	</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+biology+in+situ--the+potential+of+subtomogram+averaging.">Structural biology in situ--the potential of subtomogram averaging.</a> Curr Opin Struct Biol. 23 (2), 261-267 (2013).</li><li>Harapin, J., Eibauer, M., Medalia, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+analysis+of+supramolecular+assemblies+by+cryo-electron+tomography.">Structural analysis of supramolecular assemblies by cryo-electron tomography.</a> Structure. 21 (9), 1522-1530 (2013).</li><li>Samso, M., Wagenknecht, T., Allen, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Internal+structure+and+visualization+of+transmembrane+domains+of+the+RyR1+calcium+release+channel+by+cryo-EM.">Internal structure and visualization of transmembrane domains of the RyR1 calcium release channel by cryo-EM.</a> Nat Struct Mol Biol. 12 (6), 539-544 (2005).</li><li>Sharma, M. R., 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=Cryoelectron+microscopy+and+image+analysis+of+the+cardiac+ryanodine+receptor.">Cryoelectron microscopy and image analysis of the cardiac ryanodine receptor.</a> J Biol Chem. 273 (29), 18429-18434 (1998).</li><li>Denisov, I. G., Grinkova, Y. V., Lazarides, A. A., Sligar, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Directed+self-assembly+of+monodisperse+phospholipid+bilayer+Nanodiscs+with+controlled+size.">Directed self-assembly of monodisperse phospholipid bilayer Nanodiscs with controlled size.</a> J Am Chem Soc. 126 (11), 3477-3487 (2004).</li><li>Zhu, J., Penczek, P. A., Schroder, R., Frank, J. <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+reconstruction+with+contrast+transfer+function+correction+from+energy-filtered+cryoelectron+micrographs:+procedure+and+application+to+the+70S+Escherichia+coli+ribosome.">Three-dimensional reconstruction with contrast transfer function correction from energy-filtered cryoelectron micrographs: procedure and application to the 70S Escherichia coli ribosome.</a> J Struct Biol. 118 (3), 197-219 (1997).</li><li>Thon, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Phase contrast electron microscopy. , (1971).</li></ol>

The authors have nothing to disclose.

TEM followed by SPA has contributed many macromolecular 3D structures, nearing 2,000 entries in the EMDB by mid 2014. In general, for a given macromolecule, a low-resolution 3D structure is first determined from NS data, which may be followed by a higher-resolution cryoEM 3D structure. The high contrast of the molecular boundaries provided by NS, which facilitates the first 3D reconstruction, is later refined by cryoEM with its ability to map the internal structure of the macromolecule in its fully hydrated state, and the possibility to attain much higher resolution. Another advantage of cryoEM is the elimination of dehydration stress that could result in collapse of the macromolecule. In addition, there is the possibility to omit the carbon support, which eliminates possible surface absorption effects and shows the conformation that the macromolecule adopts in solution. Cryo-negative staining, a combination of cryoEM and NS, affords high contrast in a fully hydrated specimen and can also lead to 3D reconstruction using SPA, however it has been less frequently used, with less than 10 EMDB entries, in part because the stain itself may interfere with the integrity of the macromolecule 31. Two other TEM techniques conducive to 3D reconstruction are 2D electron crystallography and electron tomography. 2D electron crystallography requires a planar or tubular crystal; the crystal&#39;s electron diffraction is used for 3D reconstruction potentially leading to atomic resolution 32. In electron tomography of vitrified or traditionally fixed specimens, a macromolecule or sub-cellular component is rotated inside the TEM for subsequent tomographic reconstruction 33, with the advantage that singular objects can be reconstructed; however at present this technique has a resolution limit ranging from 40 to 20 &#197; 34,35. Among all these molecular TEM techniques, SPA of NS and cryoEM data has been the most widely used. The protocol illustrated here is dedicated to obtaining cryoEM images suitable for SPA analysis; nevertheless most of the protocol is also applicable to cryoEM of 2D crystals and tomographic samples.
A successful cryoEM session depends on the combined success of many critical steps; important aspects to keep in mind, explanation of common cryoEM artifacts and how to avoid them are described in the following paragraphs. This section also describes data collection guidelines to obtain high quality 3D reconstructions using the SPA method.
Avoid transition to crystalline ice. A central aspect is that the sample needs to stay in vitreous state from the moment of cryo-plunging throughout TEM observation. Thus after plunging the sample in liquid ethane all subsequent steps are performed at liquid nitrogen (-196 &#176;C) or liquid helium (-269 &#176;C) temperatures. Warming to above -135 &#176;C transforms vitreous water into crystalline water; then macromolecular rearrangements may take place and water crystals dominate the image (Figure 3A); the sample should be discarded. Accidental sample warm-up could happen if cryo-plunging, transfer of the frozen grid between containers (even if protected by a gridbox), and/or cryo holder insertion into the TEM are too slow, or if the handling tweezers were insufficiently pre-cooled. Significant vacuum deterioration in the TEM upon cryo-transfer (the cold sample traps warmer incoming air) may also warm up the sample. Finally, over-irradiation of the sample could also result in transition to crystalline ice.
Minimize ice contamination. Given the small sample volume (3-5 &#181;l) applied to the TEM grid, evaporation could concentrate buffer components (salt, detergents) and thus affect macromolecular integrity including loss of multimeric state. A high relative humidity (RH) microenvironment or chamber circumvents this problem. Alternatively cryo-plunging can be done in a sufficiently ventilated environmental cold room. After cryo-plunging RH should be as low as possible to prevent ice contamination, or condensation of ambient humidity on the cryo-grid, as the cryo-grid itself acts as a cold trap. Ice contamination consists of particles with high contrast and no substructure with sizes ranging between ~5 nm and several microns that interfere with or even totally block the image. Avoid air drafts, talking/breathing towards the cryo-grid during air transfer, and reduce ambient RH. Open liquid nitrogen containers with condensed water (visible as white suspended particles) should also be discarded.
Maximize macromolecular orientations/views. For sample support, a choice to be made is between true holey grids and thin carbon over holey grids. This depends on (i) how the sample spreads over carbon film versus the carbon holes, (ii) available sample concentration, as bare holes may require a concentration 100X higher than carbon support for a similar particle density on the image (from ~0.02 to 2 &#181;M), and (iii) how random is the distribution of macromolecule orientations. Note that glow discharge, which makes the carbon hydrophilic, will have an important effect in all three aspects and that this will be sample-dependent. Regarding the orientation of the macromolecule, while a recurrent view is desirable for the initial structural determination by random conical reconstruction, randomness of macromolecular views is desirable when refining the 3D reconstruction to higher resolutions. In our example, RyR1 interacts with the carbon with a favorite &#39;fourfold&#39; view (Figure 2D) and requires holey grids for randomness of orientations and higher resolution 36.
Maximize SNR. The best strategy to increase SNR is to reduce the background sources of noise. Excessive ice thickness and (if present) carbon film thickness beyond what is needed to support and embed the protein will add extra noise. Thus ice thickness should be reduced to the minimum necessary by controlling blotting time, pressure, RH, and filter paper quality. For carbon support, a thin (~5 nm) carbon film is layered over the thicker holey carbon film: the thick holey carbon provides mechanical resistance while the sample is imaged over the thin carbon-covered hole. On the other hand, note that extremely thin ice and/or carbon may result in an easily broken support that can turn into a web (Figure 3D). To maximize SNR, any unnecessary buffer components should be avoided. For membrane proteins, the detergent takes a significant toll on contrast (see Figure 2D, right panel). In addition by reducing surface tension the detergent changes the way in which the sample spreads on the support. Some membrane proteins require the presence of lipid in addition to detergent, which further reduces the contrast. To overcome this the sample can be diluted in a buffer with lower detergent concentration and without lipid just before cryo-plunging 37. New alternative detergents 16 and the use of nanodisks 38 to stabilize the transmembrane domain component appear to be successful approaches for imaging membrane proteins by cryoEM.
Strive for high quality images and prepare for the CTF correction. Vitrified specimens require high defocus values to generate sufficient image (phase) contrast where the CTF, the function that relates intensity to the frequency, has several zero transitions, at which point there is no information. While CTF correction is not necessary for low-resolution 3D reconstruction, when aiming for higher resolution, to recover an accurate representation of the 3D object, images with different defoci need to be collected so that computational CTF correction can be done. CTF determination and correction is included in the major SPA software packages 4-7; details can be found elsewhere 39. For optimal CTF correction, use a range of defoci. A good strategy is to alternate amongst 3-4 defocus values. The values depend on the size of the macromolecule (larger sizes require less defocus), the ice/carbon thickness (thinner sample requires less defocus), and the voltage (lower voltage requires less defocus). For 200 kV, a good starting range of values is 2.5, 3, 3.5 and 4&#160;&#181;m defocus. The CTF manifests as alternating rings of high and low intensity (Thon rings 40) in the reciprocal space representation, the power spectrum. Displaying the power spectrum will reveal the potential for resolution; the frequency of the widest visible Thon ring is the closest a priori estimate of the attainable resolution. The power spectrum should be checked periodically, and astigmatic (non circular) Thon rings (Figure 3C) should be corrected with the objective stigmator. Thon rings should be visible at least up to the resolution desired.
A cryoEM dataset obtained using this protocol can be directly processed by SPA in order to generate a 3D reconstruction. Even with an ideal sample, the quality of the 3D reconstruction will depend on the performance and specifications of the TEM equipment, and on the image processing. With the continued improvements in both these fronts, and with special mention to the recently developed direct electron detectors, the possibility to obtain 3D reconstructions at atomic resolution more consistently is closer than it has ever been.

The goal of cryoEM is to obtain electron microscopic images of macromolecules in their native hydrated state. By virtue of embedding the macromolecule in vitrified water, a frozen-hydrated sample can be directly introduced and visualized in the TEM instrument. Vitrification, achieved by flash freezing (10,000 oC/sec), freezes the sample without water crystallization and ensures that molecular rearrangements during freezing are insignificant. The excess of electron scattering of the sample with respect to the surrounding buffer provides a small but sufficient contrast to see the macromolecule in the absence of any stain 1. Computerized image processing can then be used to recreate the macromolecule&#8217;s 3D structure. Large macromolecules in the ~500 kDa- multi MDa range are ideal samples for cryoEM (representing over 80% of the Electron Microscopy Data Bank (EMDB) entries); these include proteins, protein complexes, protein-nucleic acid complexes, and membrane proteins embedded in a bilayer. These macromolecules are less likely to be crystallized (with less than 2% entries in the PDB database) yet it is often the case that smaller pieces solved by x-ray crystallography or NMR can be docked into the 3D envelope created by cryoEM. On the other hand, some of the largest structures solved by cryoEM are identifiable within the full cell by thin-section TEM 2. Thus, cryoEM effectively bridges the size and resolution gap between subcellular and atomic structures.
CryoEM reflects better the macromolecule&#8217;s native structure than the more classic method of negative staining (NS), where the macromolecule is dehydrated and surrounded by a layer of heavy metal stain whereby the region of stain exclusion creates a strongly contrasted &#8220;negative&#8220; image 3. In both cases the electron beam produces 2D projection images of the macromolecules, called particles, where the shape depends on the orientation of the macromolecule with respect to the electron beam. Many particles, at least ~1,000 and with no upper limit, can be analyzed on the computer with &#39;single particle analysis&#39; (SPA) software to increase the signal to noise ratio (SNR) though averaging, and to find the particle&#8217;s spatial orientation parameters needed to recreate the macromolecule&#8217;s 3D structure by back projection 4-7. NS, with higher SNR, is used for preliminary sample characterization and to generate a de novo 3D reconstruction; its resolution rarely exceeds 20&#160;&#197;, which is imposed by the dehydration and size of the heavy metal grain. In general, for cryoEM 3D structural determination, a low-resolution 3D reconstruction is first obtained from NS and then cryoEM is used to refine this low-resolution initial map to further study the macromolecule in its native hydrated state, to see its internal structure, and to increase its resolution. Note that if the NS model was distorted (the most common example is flattening resulting from dehydration 8) and the cryoEM particles had low SNR, then the distortion could carry into the cryoEM model (the model-bias artifact, which is common in low SNR datasets 9). Structural distortion would result in mismatch between the NS and cryoEM raw particles and between the raw cryoEM particles and corresponding projections of the 3D model orthogonal to the flattening direction. Model bias may resolve by itself as the 3D model undergoes successive refinement cycles. Should that not occur, which would be detected as lack of correspondence between raw cryoEM particles and corresponding projections from the 3D model, then a de novo model should be constructed from the cryoEM data. A good approach could be to use the angular reconstitution algorithm 10, which may yield several possible 3D volumes, and then use the NS 3D model to select the best possible cryoEM model that will be further refined 11. An even better strategy is to increase the SNR of the cryoEM dataset (see dedicated section in the Discussion).
The biochemical quality of the purified macromolecule has an utmost influence in the final outcome. The macromolecule, purified from tissue or expressed in heterologous systems, needs to have a high degree of purity and to be structurally intact. It should neither form aggregates nor should it disaggregate. To define a particular conformation, the preparation conditions should promote a uniform conformational state. To study complexes, it is optimal that their kD is in the nM range, otherwise fixatives have been successfully used to stabilize lower affinity interactions 12,13.
Unprocessed cryoEM images yield valuable information on the dimensions, general outline, state of aggregation/multimerization, homogeneity, and internal structure of the macromolecule. Nevertheless the potential of the technique is vastly enhanced with image processing. A 3D structure reveals the macromolecule&#8217;s overall architecture, 3D location of ligands and subunits, conformational changes, and enables docking of atomic structures determined by x-ray crystallography or NMR. The amount of information of a 3D reconstruction increases stepwise as the resolution is increased: in general 15-20 &#197; resolution permits unambiguous docking of atomic structures, at 9-10 &#197; alpha helices are visible as rods, at 5 &#197; it is possible to discern beta sheets, and at resolutions of 3.5 &#197; and better it is possible to build an atomic model 14.
The possibility to obtain informative images and a 3D reconstruction using SPA from relatively small quantities of sample make cryoEM an attractive technique, as 3-5 &#181;l of at least 0.05&#160;mg/ml are sufficient to prepare one TEM grid. The minimum instrumental requirements for cryoEM are a home-made or commercial cryo-plunger, an electron microscope with ultra high vacuum, image recording media and low-dose kit, a cryo-holder with its pumping station, a glow discharge apparatus, and an evaporator. Non-essential but further desirable features on the TEM are CCD camera (present in all modern TEMs) or a direct electron detector, field emission gun (FEG), energy filtering, and high throughput data collection software. This software in principle enables collecting tens of thousands of particles in a single cryoEM session 15, however the increased data storage needs and subsequent supervised post-processing time need to be taken into consideration. FEG combined with contrast transfer function (CTF) correction underlies most 3D reconstructions that reached resolution sufficient to visualize alpha helices. At that point, the level of resolution is also sample-dependent: reaching 10&#160;&#197; resolution is less common for membrane proteins whereas if high order of symmetry is present then atomic resolution is more achievable. The recent development of direct electron detectors has enabled atomic resolution even for macromolecules with low (4x) or no symmetry, where the quality of the cryoEM electron density maps matches these derived from x-ray diffraction data 16,17.
Practical examples illustrating the capabilities of the technique are provided here for four important types of macromolecules where cryoEM has a continuing chief role in their structural determination. (i) With their icosahedral symmetry that increases the dataset by 60-fold and their large size that permits faithful and reproducible alignment, the structures of several icosahedral viruses have been resolved to near-atomic resolution by cryoEM followed by SPA 18. We show an example of a class of icosahedral viruses, the adeno-associated viruses (AAVs), for which near-atomic structures by cryoEM and by cryoEM/x-ray hybrid methods exist 19,20. (ii) Macromolecules with helical structure include microtubules, filaments and viruses. Their repetitive units along the helical axis can be analyzed by a more complex version of SPA adapted to the helical geometry 21. In the example we show tobacco mosaic virus (TMV), an RNA virus that has been solved to atomic resolution by cryoEM 22. (iii) Large soluble proteins have been amply studied. Among these, macromolecules forming symmetrical multimeric cylinders constitute a recurring architecture often solved at subnanometer resolution 23,24. Here we show images of KLH, an invertebrate oxygen carrier, where a hybrid molecular model was created from cryoEM/x-ray crystallography hybrid methods 25. (iv) Membrane proteins are an important class of proteins; they constitute about 1/3 of the proteins coded by the human genome, yet they are difficult to characterize structurally due to complexities associated with transmembrane domain stabilization 26, constituting less than 1.5% of the structures solved by any structural technique combined. The role of cryoEM and 3D reconstruction in their structural determination is exemplified with the ryanodine receptor (RyR), a large eukaryotic intracellular Ca2+ channel important in muscle contraction and brain signaling. The highest resolution cryoEM structures to date, with 10 &#197; resolution, reveal secondary structure 27.

<table border="0" cellpadding="0" cellspacing="0" width="661">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Name of Material/ Equipment</td>
			<td style="width:132px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="120">
			<td height="120" style="height:120px;width:243px;">Ethane</td>
			<td style="width:132px;">Airgas</td>
			<td align="right" style="width:119px;">371745</td>
			<td style="width:167px;">99.99% purity very important, impurities can lead to high-contrast contaminants on the grid. CAUTION: flammable</td>
		</tr>
		<tr height="160">
			<td height="160" style="height:160px;width:243px;">Liquid nitrogen</td>
			<td style="width:132px;">Airgas</td>
			<td align="right" style="width:119px;">27135</td>
			<td style="width:167px;">99% purity. CAUTION: handling liquid nitrogen in a cold room poses a serious asphyxia hazard since nitrogen displaces oxygen without any warning signs.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dumont no. 5 Antimagnetic tweezers</td>
			<td style="width:132px;">Ted Pella</td>
			<td align="right" style="width:119px;">5326</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Mica sheets, 1 x 4 cm</td>
			<td style="width:132px;">Ted Pella</td>
			<td align="right" style="width:119px;">54</td>
			<td></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:243px;">Graphite rods, presharpened size 1/8&#8221; dia x 2 &#188;&#8221; long tip 0.039&#8221; dia neck type</td>
			<td style="width:132px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">70220-45</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:243px;">350 ml cryo-dewars</td>
			<td style="width:132px;">Pope</td>
			<td align="right" style="width:119px;">8600</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:243px;">Cu 400 mesh holey grids&#160; Quantifoil</td>
			<td style="width:132px;">Quantifoil</td>
			<td style="width:119px;">Q10525</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:243px;">Cu 400 mesh holey grids C-Flat</td>
			<td style="width:132px;">Protochips</td>
			<td style="width:119px;">CF-1.2/1.3-4C</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:243px;">Glow discharge device</td>
			<td style="width:132px;">Electron Microscopy Sciences</td>
			<td style="width:119px;">Model E7620</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:243px;">Turbo pumping station</td>
			<td style="width:132px;">Gatan</td>
			<td style="width:119px;">Model 655</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:243px;">Carbon evaporator</td>
			<td style="width:132px;">Denton Vaccum</td>
			<td style="width:119px;">Model 502B</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:243px;">Freeze plunger</td>
			<td style="width:132px;">FEI</td>
			<td style="width:119px;">Vitrobot Mark IV</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:243px;">Image Processing software CTFFIND3/CTFTILT</td>
			<td style="width:132px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">http://grigoriefflab.janelia.org/ctf</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:243px;">Image Processing software EMAN</td>
			<td style="width:132px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">http://blake.bcm.edu/emanwiki/EMAN2</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;width:243px;">Image Processing software Spider</td>
			<td style="width:132px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">http://spider.wadsworth.org/spider_doc/spider/docs/spider.html</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;width:243px;">Image Processing software Freealign&#160;</td>
			<td style="width:132px;"></td>
			<td style="width:119px;"></td>
			<td style="width:167px;">http://grigoriefflab.janelia.org/frealign</td>
		</tr>
		<tr height="41">
			<td height="41" style="height:41px;">3D visualization software Chimera</td>
			<td></td>
			<td></td>
			<td style="width:167px;">http://www.cgl.ucsf.edu/chimera/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Native Keyhole Limpet Hemocyanin</td>
			<td style="width:132px;">Biosyn</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

do's and don'ts of cryo-electron microscopy: a primer on sample preparation and high quality data collection for macromolecular 3d reconstruction

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state by transmission electron microscopy (TEM). This can be achieved with very low quantity of protein and in the buffer of choice, without the use of any stain, which is very useful to determine structure-function correlations of macromolecules. When combined with single-particle image processing, the technique has found widespread usefulness for 3D structural determination of purified macromolecules. 
The protocol presented here explains how to perform cryoEM and examines the causes of most commonly encountered problems for rational troubleshooting; following all these steps should lead to acquisition of high quality cryoEM images. The technique requires access to the electron microscope instrument and to a vitrification device. Knowledge of the 3D reconstruction concepts and software is also needed for computerized image processing. Importantly, high quality results depend on finding the right purification conditions leading to a uniform population of structurally intact macromolecules. 
The ability of cryoEM to visualize macromolecules combined with the versatility of single particle image processing has proven very successful for structural determination of large proteins and macromolecular machines in their near-native state, identification of their multiple components by 3D difference mapping, and creation of pseudo-atomic structures by docking of x-ray structures. The relentless development of cryoEM instrumentation and image processing techniques for the last 30 years has resulted in the possibility to generate de novo 3D reconstructions at atomic resolution level.

Careful execution of all the protocol steps outlined in Figure 1 enables visualization of frozen hydrated, non-stained macromolecules. Results for four representative specimens with different structural characteristics are shown. Their microscopic images obtained by the methods of NS and cryoEM are compared (Figure 2). (i) NS images of AAV5 reveal virus-like particles of around 130 &#197; diameter. The coexistence of full and empty structures corresponds to broken (that allow stain entry) and structurally intact capsids, respectively. CryoEM shows uniformly empty structures; indeed these viral particles do not contain genetic material 28. NS shows predominant round particles whereas cryoEM shows mostly hexagonal particles, reflecting their icosahedral form. (ii) Images of the helical TMV show rods of 180 &#197; diameter and variable length, often longer than 0.1 &#181;m, with a helical repeat of 23 &#197; visible both in NS and in cryoEM. The 40 &#197; central channel is filled with stain in the NS images and empty in the cryoEM images. (iii) Native KLH, a didecamer of 8 MDa, assembles in characteristic barrels of 350 &#197; diameter and 400 &#197; length. NS and cryoEM reveal KLH&#8217;s rectangular side views and circular end-on views. Both techniques reveal six striations perpendicular to the symmetry axis and the equally spaced morphological units within each tier. CryoEM reveals an empty inner structure. (iv) Ryanodine receptor, a large tetrameric intracellular channel of 2.26 MDa, is seen as a square of 275&#160;x 275&#160;&#197;2. Intricate surface features such as a central cross and a central depression manifest as stain accumulation by NS, and as lower density regions by cryoEM. While NS shows similar contrast in all four tested samples, comparison of the cryoEM panels exposes the lower SNR (contrast) of RyR1 due to the presence of detergent to solubilize this membrane protein.
Typical examples of cryoEM artifacts are shown for RyR1: crystalline ice, ice contamination, astigmatism, and web-like structures (Figure 3). Their causes and prevention are further described in the discussion section.
SPA and 3D reconstruction of frozen-hydrated RyR1 reveals its fourfold symmetric structure, which reflects the protein&#8217;s homotetrameric organization (Figure 4A). The cytoplasmic domain forms a square prism of 275 x 275 x 120 &#197;3 and contains several reproducible globular domains forming a scaffold-like structure. The transmembrane domain forms a smaller tapered square prism with maximum dimensions 115 x 115 x 60 &#197;3. At 10 &#197; resolution it is possible to visualize alpha helices (Figure 4B). 3D difference mapping can reveal the position of important ligands, in this case the 3D difference map of FKBP12, a 12 kD protein considered a subunit of RyR1, is superimposed on RyR1 (Figure 4C) 29. Finally, conformational changes provide a dynamic view of the macromolecule. In this case RyR1 previously stabilized in either closed or open conditions reveals a mass translocation from the closed to the open state (Figure 4D) 27.
<img alt="Figure 1" src="/files/ftp_upload/52311/52311fig1.jpg" /> 
Figure 1. Flow diagram of the cryo electron microscopy technique. The diagram highlights the main steps of the protocol.
<img alt="Figure 2" src="/files/ftp_upload/52311/52311fig2.jpg" /> 
Figure 2. Comparison of NS and cryoEM for a variety of samples. (A) AAV5, an icosahedral virus. (B) TMV, a helical virus. Insets show the helical repeat, of 23 &#197;. (C) Native KLH. (D) RyR1; representative views are highlighted (f, fourfold view and s, side view). All samples are imaged under low dose conditions and at the same magnification of 50,000X an under an accelerating voltage of 200 kV. Scale bars, 10 nm. All NS samples are on carbon support, cryoEM samples A, C, D are on thin carbon over quantifoil, and cryoEM sample B is directly over quantifoil holes. <a href="https://www.jove.com/files/ftp_upload/52311/52311fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52311/52311fig3.jpg" /> 
Figure 3. Gallery of images showing common cryoEM artifacts on an RyR1 cryoEM sample. (A) Crystalline ice. (B) Ice contamination (arrows). (C) Astigmatism. RyR1s in the astigmatic image are barely visible. The inset on the right shows the power spectrum of the image with asymmetric Thon rings. The inset on the left shows the power spectrum of a non-astigmatic image for reference. (D) Web-like structures. <a href="https://www.jove.com/files/ftp_upload/52311/52311fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52311/52311fig4.jpg" /> 
Figure 4. CryoEM and 3D reconstruction of the ryanodine receptor. (A) Isosurface representation. (B) Docking of a crystal structure of RyR1&#8217;s N-terminus 30 showing good agreement between the atomic coordinates and the cryoEM envelope; arrows point to two alpha helices that protrude from the structure. Arrowhead indicates one of the four helices that surround the pore of the ion channel. The dotted lines indicate the boundaries of the membrane. (C) RyR1 and 3D location of the FKBP12 ligand (orange color). (D) Conformational changes of RyR1 in going from the closed (orange) to the open (black mesh) conformation. <a href="https://www.jove.com/files/ftp_upload/52311/52311fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Do's and Don'ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction at JoVE.com

Do&#039;s and Don&#039;ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction

This paper describes the cryo-electron microscopy methodology, which is used to obtain high quality microscopic images of macromolecules in their near-native state. The method yields images suitable for further computerized processing using the single particle approach, devised to generate the 3D reconstruction of a macromolecule.

Do's and Don'ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction

Cryo-electron microscopy (cryoEM) entails flash-freezing a thin layer of sample on a support, and then visualizing the sample in its frozen hydrated state ...

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Biology

45.9K Views.  Virginia Commonwealth University. This paper describes the cryo-electron microscopy methodology, which is used to obtain high quality microscopic images of macromolecules in their near-native state. The method yields images suitable for further computerized processing using the single particle approach, devised to generate the 3D reconstruction of a macromolecule.

Video: Do's and Don'ts of Cryo-electron Microscopy: A Primer on Sample Preparation and High Quality Data Collection for Macromolecular 3D Reconstruction

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Preparation of Quality Inositol Pyrophosphates

Myo-inositol is present in nature either unmodified or in more complex phosphorylated derivates. Of the latest, the two most abundant in eukaryotic cells are inositol pentakisphosphate (IP5) and inositol hexakisphosphate (phytic acid or IP6). IP5 and IP6 are the precursors of inositol pyrophosphate molecules that contain one or more pyrophosphate bonds1. Phosphorylation of IP6 generates diphoshoinositolpentakisphosphate (IP7 or PP-IP5) and bisdiphoshoinositoltetrakisphosphate (IP8 or (PP)2-IP4). Inositol pyrophosphates have been isolated from all eukaryotic organisms so far studied. In addition, the two distinct classes of enzymes responsible for inositol pyrophosphate synthesis are highly conserved throughout evolution2-4.
The IP6 kinases (IP6Ks) posses an enormous catalytic flexibility, converting IP5 and IP6 to PP-IP4 and IP7 respectively and subsequently, by using these products as substrates, promote the generation of more complex molecules5,6. Recently, a second class of pyrophosphate generating enzymes was identified in the form of the yeast protein VIP1 (also referred as PP-IP5K), which is able to convert IP6 to IP7 and IP87,8.
Inositol pyrophosphates regulate many disparate cellular processes such as insulin secretion9, telomere length10,11, chemotaxis12, vesicular trafficking13, phosphate homeostasis14 and HIV-1 gag release15. Two mechanisms of actions have been proposed for this class of molecules. They can affect cellular function by allosterically interacting with specific proteins like AKT16. Alternatively, the pyrophosphate group can donate a phosphate to pre-phosphorylated proteins17. The enormous potential of this research field is hampered by the absence of a commercial source of inositol pyrophosphates, which is preventing many scientists from studying these molecules and this new post-translational modification. The methods currently available to isolate inositol pyrophosphates require sophisticated chromatographic apparatus18,19. These procedures use acidic conditions that might lead to inositol pyrophosphate degradation20 and thus to poor recovery. Furthermore, the cumbersome post-column desalting procedures restrict their use to specialized laboratories.
In this study we describe an undemanding method for the generation, isolation and purification of the products of the IP6-kinase and PP-IP5-kinases reactions. This method was possible by the ability of polyacrylamide gel electrophoresis (PAGE) to resolve highly phosphorylated inositol polyphosphates20. Following IP6K1 and PP-IP5K enzymatic reactions using IP6 as the substrate, PAGE was used to separate the generated inositol pyrophosphates that were subsequently eluted in water.

Inositol pyrophosphates play an important role in human pathologies such cancer, diabetes and obesity; however, the exact mechanism of action is a matter of dispute. The lack of commercially available inositol pyrophosphates renders detailed studies problematic. Here we describe a simple protocol to produce and isolate milligrams of inositol pyrophosphates.

Myo-inositol is present in nature either unmodified or in more complex phosphorylated derivates. Of the latest, the two most abundant in eukaryotic cells ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Cryo-electron Microscopy Specimen Preparation By Means Of a Focused Ion Beam

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms or solutions. We make use of a custom built cryo-transfer station and a modified cryo-SEM preparation chamber2. The spores are taken from a culture, plunge-frozen in a liquid nitrogen slush and observed in the cryo-SEM to select a region of interest. A thin lamella is then extracted using the FIB, attached to a TEM grid and subsequently thinned to electron transparency. The grid is transferred to a cryo-TEM holder and into a TEM for high resolution studies. Thanks to the introduction of a cooled nanomanipulator tip and a cryo-transfer station, this protocol is a straightforward adaptation to cryogenic temperature of the routinely used FIB preparation of TEM samples. As such it has the advantages of requiring a small amount of modifications to existing instruments, setups and procedures; it is easy to implement; it has a broad range of applications, in principle the same as for cryo-TEM sample preparation. One limitation is that it requires skillful handling of the specimens at critical steps to avoid or minimize contaminations.

Cryo Electron Microscopes, either Scanning (SEM) or Transmission (TEM), are widely used for characterization of biological samples or other materials with a high water content1. A SEM/Focused Ion Beam (FIB) is used to identify features of interest in samples and extract a thin, electron-transparent lamella for transfer to a cryo-TEM.

Here we present a protocol used to prepare cryo-TEM samples of Aspergillus niger spores, but which can easily be adapted for any number of microorganisms ...

Sample Preparation and Imaging of Exosomes by Transmission Electron Microscopy

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The contents and morphology of the secreted vesicles reflect cell behavior or physiological status, for example cell growth, migration, cleavage, and death. The exosomes&#39; role may depend highly on size, and the size of exosomes varies from 30 to 300 nm. The most widely used method for exosome imaging is negative staining, while other results are based on Cryo-Transmission Electron Microscopy, Scanning Electron Microscopy, and Atomic Force Microscopy. The typical exosome&#39;s morphology assessed through negative staining is a cup-shape, but further details are not yet clear. An exosome well-characterized through structural study is necessary particular in medical and pharmaceutical fields. Therefore, function-dependent morphology should be verified by electron microscopy techniques such as labeling a specific protein in the detailed structure of exosome. To observe detailed structure, ultrathin sectioned images and negative stained images of exosomes were compared. In this protocol, we suggest transmission electron microscopy for the imaging of exosomes including negative staining, whole mount immuno-staining, block preparation, thin section, and immuno-gold labelling.

This protocol describes the various techniques necessary for transmission electron microscopy including negative staining, ultrathin sectioning for detailed structure, and immuno-gold labelling to determine the positions of specific proteins in exosomes.

Exosomes are nano-sized extracellular vesicles secreted by body fluids and are known to represent the characteristics of cells that secrete them. The ...

Visualization of DNA Compaction in Cyanobacteria by High-voltage Cryo-electron Tomography

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to occur in some cyanobacteria before cell division. However, due to the large cell size and the transient character, it is difficult to investigate the structure in detail. To overcome the difficulties, first, DNA compaction is reproducibly produced in the cyanobacterium Synechococcus elongatus PCC 7942 by synchronous culture using 12 h each light/dark cycle. Second, DNA compaction is monitored by fluorescence microscopy and captured by rapid freezing. Third, the detailed structure of DNA compacted cells is visualized in three dimensions (3D) by high voltage cryo-electron tomography. This set of methods is widely applicable to investigate transient structures in bacteria, e.g. cell division, chromosome segregation, phage infection etc., which are monitored by fluorescence microscopy and directly visualized by cryo-electron tomography at appropriate time points.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. Synchronous cultivation, monitoring by fluorescence microscopy, rapid freezing, and high voltage cryo-electron tomography are used. A protocol for these methodologies is presented, and future applications and developments are discussed.

This protocol describes how to visualize the transient DNA compaction in cyanobacteria. DNA compaction is a dramatic cytoplasmic event recently found to ...

Quantification of Site-specific Protein Lysine Acetylation and Succinylation Stoichiometry Using Data-independent Acquisition Mass Spectrometry

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein functions, for example, by changing enzymatic activity or by mediating interactions. Precise quantification of site-specific protein acylation occupancy, or stoichiometry, is essential for understanding the functional consequences of both global low-level stoichiometry and individual high-level acylation stoichiometry of specific lysine residues. Other groups have reported measurement of lysine acetylation stoichiometry by comparing the ratio of peptide precursor isotopes from endogenous, natural abundance acylation and exogenous, heavy isotope-labeled acylation introduced after quantitative chemical acetylation of proteins using stable isotope-labeled acetic anhydride. This protocol describes an optimized approach featuring several improvements, including: (1) increased chemical acylation efficiency, (2) the ability to measure protein succinylation in addition to acetylation, and (3) improved quantitative accuracy due to reduced interferences using fragment ion quantification from data-independent acquisitions (DIA) instead of precursor ion signal from data-dependent acquisition (DDA). The use of extracted peak areas from fragment ions for quantification also uniquely enables differentiation of site-level acylation stoichiometry from proteolytic peptides containing more than one lysine residue, which is not possible using precursor ion signals for quantification. Data visualization in Skyline, an open source quantitative proteomics environment, allows for convenient data inspection and review. Together, this workflow offers unbiased, precise, and accurate quantification of site-specific lysine acetylation and succinylation occupancy of an entire proteome, which may reveal and prioritize biologically relevant acylation sites.

Here, we present unbiased quantification of site-specific protein acetylation and/or succinylation occupancy (stoichiometry) of an entire proteome through a ratiometric analysis of endogenous modifications to modifications introduced after quantitative chemical acylation using stable isotope-labeled anhydrides. In combination with sensitive data-independent acquisition mass spectrometry, accurate site occupancy measurements are obtained.

Post-translational modification (PTM) of protein lysine residues by N&#400;-acylation induces structural changes that can dynamically regulate protein ...

Miniaturized Sample Preparation for Transmission Electron Microscopy

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein complexes to atomic resolution. However, protein isolation techniques and sample preparation methods for EM remain a bottleneck. A relatively small number (100,000 to a few million) of individual protein particles need to be imaged for the high-resolution analysis of proteins by the single particle EM approach, making miniaturized sample handling techniques and microfluidic principles feasible.
A miniaturized, paper-blotting-free EM grid preparation method for sample pre-conditioning, EM grid priming and post processing that only consumes nanoliter-volumes of sample is presented. The method uses a dispensing system with sub-nanoliter precision to control liquid uptake and EM grid priming, a platform to control the grid temperature thereby determining the relative humidity above the EM grid, and a pick-and-plunge-mechanism for sample vitrification. For cryo-EM, an EM grid is placed on the temperature-controlled stage and the sample is aspirated into a capillary. The capillary tip is positioned in proximity to the grid surface, the grid is loaded with the sample and excess is re-aspirated into the microcapillary. Subsequently, the sample film is stabilized and slightly thinned by controlled water evaporation regulated by the offset of the platform temperature relative to the dew-point. At a given point the pick-and-plunge mechanism is triggered, rapidly transferring the primed EM grid into liquid ethane for sample vitrification. Alternatively, sample-conditioning methods are available to prepare nanoliter-sized sample volumes for negative stain (NS) EM.
The methodologies greatly reduce sample consumption and avoid approaches potentially harmful to proteins, such as the filter paper blotting used in conventional methods. Furthermore, the minuscule amount of sample required allows novel experimental strategies, such as fast sample conditioning, combination with single-cell lysis for &#34;visual proteomics,&#34; or &#34;lossless&#34; total sample preparation for quantitative analysis of complex samples.

An instrument and methods for the preparation of nanoliter-sized sample volumes for transmission electron microscopy is presented. No paper-blotting steps are required, thus avoiding the detrimental consequences this can have for proteins, significantly reducing sample loss and enabling the analysis of single cell lysate for visual proteomics.

Due to recent technological progress, cryo-electron microscopy (cryo-EM) is rapidly becoming a standard method for the structural analysis of protein ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.