We would like to thank Peter A. Kahn and Terry Rohde at Engineering Arts LLC (Arizona, USA) for initial design and subsequent development of the Spotiton system. We thank the staff of the Simons Electron Microscopy Center at the New York Structural Biology Center for help and technical support. The work presented here was conducted at the National Resource for Automated Molecular Microscopy located at the New York Structural Biology Center, supported by grants from the NIH (GM103310) and the Simons Foundation (SF349247).

1. Set up the Spotiton machine and software
<ol>
	<li>Prepare the system to dispense (Figure 1).
	<ol>
		<li>Open the main valve on the nitrogen supply tank. Ensure the system reservoir is filled with degassed, ultrapure water. Turn on the computer. Turn on the Spotiton system at the multioutlet powerstrip.</li>
		<li>Click on the desktop icon (Figure 2A) to open the user interface (UI) of the Spotiton software (Figure 2B). In the Tools menu, select Initialize Stages to initialize and home the 3-axis robots (grid stage and pipette stage) and the rotating dispenser head assembly (theta stage). Ensure the dispenser tips are pointing down prior to initialization and homing.</li>
		<li>On the main window, click on Go to SafePosition to send the robots to the SafePosition (Figure 3). 
		NOTE: Moving to SafePosition can be done with robots in any position without risk of collision.</li>
		<li>On the Aspirate tab, select Prime to flush the dispenser heads multiple times with water from the reservoir. Continue until&#160; uninterrupted streams of water can be seen emerging from the two tips. 
		NOTE: This may need to be repeated if a significant volume of air entered the fluid lines when the tips were flushed with methanol at last shutdown.</li>
	</ol>
	</li>
	<li>Inspect dispenser performance
	<ol>
		<li>On the Inspect tab, send each tip to the inspection camera to test-fire water, and match the pattern of droplet production from the two dispensers (Figure S1). 
		NOTE: If a tip fails to fire due to a bubble (Figure S2A) or debris, clean the tips at the wash station (Figure S2B) using the Ultrasonic Wash function accessed on the Aspirate tab.</li>
		<li>Adjust the firing amplitude (unitless) for each dispenser and sample to achieve a stream of discrete droplets of consistent size.</li>
		<li>Visually confirm equivalent droplet production by the two dispensers.
		<ol>
			<li>Press the Record button in the upper camera monitor to record a video of Tip 1 firing. Play back the video of Tip 1 firing in the right-side monitor at the same time Tip 2 is fired in the upper camera monitor (Figure S1). 
			NOTE: Videos of each tip firing are recorded and stored.</li>
		</ol>
		</li>
		<li>Dispensers are now ready to test-fire on a grid. 
		NOTE: This protocol assumes that correct alignment of the grid and pipette stages at their respective plunge positions has been verified. Failure to confirm correct alignment could result in a collision, damaging the tips or the robots.</li>
	</ol>
	</li>
	<li>Test-fire water on a grid.
	<ol>
		<li>Ensure robots are at the SafePosition.</li>
		<li>Remove the tweezers from the mount on the grid robot using the supplied Allen key.</li>
		<li>On a nearby benchtop, position a test grid, nanowire side up, on the edge of the grid block. Carefully grab the rim of the grid, position correctly in the tweezers (Figure S3), and remount the tweezers.</li>
		<li>On the Cryo tab, click on Tip to Camera to move the tips into the field of view of the upper plunge path camera (upper camera). Ensure Live is selected in the upper camera monitor, and the upper camera light is turned on (dial on front of machine cabinet (Figure 1).</li>
		<li>Mount the tweezers on the grid robot. Position Tip 1, visible in the upper camera monitor, by clicking the mouse within the monitor. 
		NOTE: Only Tip 1 will be visible and will move to the location of the click.</li>
		<li>Click Grid to Camera to position the grid in front of the upper camera. Adjust Tip 1 position as before, if needed.</li>
		<li>Choose either Tip1, Tip2, or Tip1 and Tip2 to select either one or both dispensers to test-fire. 
		NOTE: When test-firing the tips individually, use the mouse to position Tip 1 in the upper camera monitor at different lateral locations on the grid. This will deposit liquid stripes from the two tips in a non-overlapping pattern and allow the user to confirm each tip has fired.</li>
		<li>On the Cryo tab, ensure Vitrify Grid is not selected, click on Queue Target, then on Plunge. 
		NOTE: The pipette stage will rise slightly, followed by the grid stage. The grid robot then plunges the grid past the firing dispensers and upper and lower cameras, coming to rest just above the deck. An image capture from the upper camera appears above an image capture from the lower camera on the left-hand side of the UI.</li>
		<li>Evaluate the upper and lower images: Did each tip fire when selected? When fired together, did the liquid from the two tips overlap fully, creating a noticeably thicker stripe than when fired individually?
		<ol>
			<li>If either tip failed to fire, perform one or multiple ultrasonic wash cycles until clear firing is observed.</li>
			<li>If the sample stripes do not fully overlap, adjust the lateral alignment of one of the dispensers using an Allen key to turn the lateral adjustment screw on one of the dispensers one quarter turn, and perform a test plunge. Repeat until stripes fully overlap. 
			&#8203;NOTE: If both tips fired successfully and as expected, the system is ready to prepare sample grids.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Prepare two-sample, mixed grids
<ol>
	<li>Update the acceleration, deceleration, and velocity values in the Machine Parameters XML file to attain the mixing time of interest. 
	NOTE: Tables correlating these values to mixing times have been previously reported15.</li>
	<li>Ready the sample and grids. 
	NOTE: From this point in the protocol, 20-30 min will elapse before the first grid is prepared, and samples should be kept at an appropriate temperature during this time interval.
	<ol>
		<li>Dilute two samples (i.e., protein and ligand or other interacting partner) to the desired concentrations with an appropriate buffer, ideally, the same for both. 
		NOTE: Spotiton grids require 1.5-2 times higher concentrations of protein than typically used for automatic plunge freezers. This may be due to the minimal time the protein spends in a thin aqueous layer on the grid surface during a plunge, giving particles less opportunity to concentrate at the air-water interface.</li>
		<li>Fill the cryogen bowl with liquid nitrogen.</li>
		<li>Plasma clean 3-4 nanowire grids. Use 5 W, hydrogen and oxygen, 1.5 min as a starting point. 
		NOTE: The most effective plasma cleaning duration and recipe may change from day-to-day based on ambient temperature and humidity and the batch of nanowire grids used. Alternative gas mixtures or a glow discharger may also be effective, but have not been tested for this purpose. As water and protein in buffer solution are often wicked at different speeds by nanowires, it is best practice to evaluate the wicking of the grids to be used that day on a plunge in which the actual sample is dispensed.</li>
	</ol>
	</li>
	<li>Set the humidity level in the system. 
	NOTE: In addition to the conditions used to both make and plasma clean the grids, humidity is another primary factor that impacts the wicking speed of nanowire grids. Although the target percent humidity for a particular session is determined empirically for each day and batch of grids, 90-95% is a good starting point.
	<ol>
		<li>Ensure the nebulizer is filled sufficiently with ultrapure water. Plug in the nebulizer, and observe the vapor exit the central port on the nebulizer cap.</li>
		<li>Observe the live humidity monitor in the main window, or open the ambient humidity tracker under Reports | Ambient (Figure S4). Check humidity levels in the two zones: Chamber and Shroud. 
		NOTE: Chamber includes all areas within the Spotiton enclosure. Shroud is the area immediately encompassing the &#34;at camera&#34; positions of the grid and dispenser tips. The black resin shroud maintains the set humidity level when the enclosure doors are opened to load a grid.</li>
		<li>Once the target humidity values are reached, maintain these values either automatically or manually from the Humidity tab. Select Automatic control, and choose a setpoint and threshold to maintain the setpoint within the threshold limit chosen. Alternatively, select Manual control, and adjust the humidity level using the two fan controls: chamber fan and shroud fan. 
		NOTE: Turning on the chamber fan pulls the vapor through a filter in the left port and prevents larger water droplets from entering the chamber, preventing further increase in ambient humidity. As long as the chamber doors remain closed, the humidity level will stabilize at the desired percentage. Turning on the shroud fan pulls the vapor through the right port into the shroud. This both reduces vapor release into the chamber and increases the humidity level within the shroud.</li>
	</ol>
	</li>
	<li>Load the sample into the dispensers.
	<ol>
		<li>Add 5 &#181;L of each sample into the sample cups. 
		NOTE: To avoid introducing bubbles, dispense the volume very carefully onto the inner sidewall of the cup, then shake down to force the sample to the bottom of the cup.</li>
		<li>Load the sample cups into the holding tray, the sample for Tip 1 on the left, for Tip 2 on the right. Push the tray back into the machine until it seats.</li>
		<li>On the Aspirate tab, select 3 &#181;L for the volume to be aspirated by each tip. Ensure the sample tray has been seated securely, then click on Aspirate and observe how the pipette stage moves the dispenser heads into the sample cups.</li>
		<li>Verify successful aspiration of both samples by removing the sample cups and observing a drop in the liquid level.</li>
		<li>On the Inspect tab, send each tip to the inspection camera to confirm unobstructed dispensing. Adjust the amplitude as needed to match droplet formation from each tip (see section 1.2). 
		NOTE: Amplitude will likely need to be increased for the tip that dispenses the protein sample.</li>
		<li>The system is now ready to prepare a sample grid.</li>
	</ol>
	</li>
	<li>Freeze sample grids
	<ol>
		<li>Load a freshly plasma cleaned grid into the tweezers, but do not mount the tweezers yet. Ensure humidity level is elevated, ~90-95%. Fill the ethane cup.</li>
		<li>Perform a final test-fire of both tips in front of the inspection camera, confirming no obstruction. On the Cryo tab, click on Tip to camera.</li>
		<li>Fill ethane cup. If ethane ice forms, melt as needed with additional ethane gas. 
		NOTE: Steps 2.5.4 and 2.5.5 should be completed relatively quickly (&#60;20 s) to minimize (i) saturation of the nanowires in the high humidity of the chamber and (ii) the likelihood that the tip firing the protein sample will clog.</li>
		<li>Mount the tweezers with the grid onto the grid stage. On the Cryo tab, click on Grid to camera. Ensure Tip 1 is positioned correctly in the upper camera monitor (see steps 1.3.4-1.3.5).</li>
		<li>Click on Vitrify Grid, Queue Target, then Plunge. Click on OK when prompted to command the grid robot to hop the grid from ethane into liquid nitrogen and release it onto the submerged shelf. 
		NOTE: The tweezers then rise back into the chamber. After the hop to liquid nitrogen, a prompt appears asking if the grid dropped from the tweezers. If it did not drop, click on No, and the tweezers will open and close several times to detach the grid.</li>
		<li>Examine images of the grid (Figure S5) to decide if it should be kept or discarded.</li>
		<li>If keeping the grid, transfer it to the pre-cooled grid box. Alternatively, spot subsequent grids and then transfer all grids at once to the grid box, taking care that each grid can be identified by its position in the resting area.</li>
		<li>To transfer the grid to a grid box, pre-cool fine-tipped forceps, gently grasp the grid by the edge, and place it into a grid box slot, starting with the first slot left of the notch going clockwise.</li>
		<li>When all the grids are loaded, attach the grid box lid to the lid tool, and precool in liquid nitrogen. Screw the lid onto the grid box, and tighten with a lid-tool or a precooled screwdriver.</li>
		<li>Use large forceps to quickly transfer the closed grid box to a small dewar for imaging or long-term storage.</li>
	</ol>
	</li>
	<li>Evaluate grid suitability for downstream EM imaging.
	<ol>
		<li>Observe the images from the upper and lower plunge path cameras that are displayed on the left side of the UI immediately after a grid is plunged. Use these images to evaluate wicking speed, successful firing of both tips, and the extent of overlap of the two samples stripes.</li>
		<li>Review and compare the grid images from the upper and lower cameras along with the machine settings and humidity measurements at the time of the plunge using the experiment viewer: Reports | Experiment.</li>
		<li>Select grids for imaging in the electron microscope that demonstrate evidence of good wicking.</li>
	</ol>
	</li>
</ol>

<ol><li>Berriman, J., Unwin, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Analysis+of+transient+structures+by+cryo-microscopy+combined+with+rapid+mixing+of+spray+droplets%5BTitle%5D)+AND+%22Ultramicroscopy%22%5BJournal%5D)">Analysis of transient structures by cryo-microscopy combined with rapid mixing of spray droplets.</a> Ultramicroscopy. 56 (4), 241-252 (1994).</li>
<li>Dubochet, J., McDowall, A. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Vitrification+of+pure+water+for+electron+microscopy%5BTitle%5D)+AND+%22Journal+of+microscopy%22%5BJournal%5D)">Vitrification of pure water for electron microscopy.</a> Journal of microscopy. 124 (3), 3-4 (1981).</li>
<li>White, H. D., Walker, M. L., Trinick, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+computer-controlled+spraying-freezing+apparatus+for+millisecond+time-resolution+electron+cryomicroscopy%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">A computer-controlled spraying-freezing apparatus for millisecond time-resolution electron cryomicroscopy.</a> Journal of Structural Biology. 121 (3), 306-313 (1998).</li>
<li>White, H. D., Thirumurugan, K., Walker, M. L., Trinick, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+second+generation+apparatus+for+time-resolved+electron+cryo-microscopy+using+stepper+motors+and+electrospray%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">A second generation apparatus for time-resolved electron cryo-microscopy using stepper motors and electrospray.</a> Journal of Structural Biology. 144 (1-2), 246-252 (2003).</li>
<li>Kontziampasis, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+cryo-EM+grid+preparation+device+for+time-resolved+structural+studies%5BTitle%5D)+AND+%22IUCrJ%22%5BJournal%5D)">A cryo-EM grid preparation device for time-resolved structural studies.</a> IUCrJ. 6, Pt 6 1024-1031 (2019).</li>
<li>Lu, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Gas-assisted+annular+microsprayer+for+sample+preparation+for+time-resolved+cryo-electron+microscopy%5BTitle%5D)+AND+%22Journal+of+Micromechanics+and+Microengineering%3A+Structures%2C+Devices%2C+and+Systems%22%5BJournal%5D)">Gas-assisted annular microsprayer for sample preparation for time-resolved cryo-electron microscopy.</a> Journal of Micromechanics and Microengineering: Structures, Devices, and Systems. 24 (11), 115001(2014).</li>
<li>Chen, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structural+dynamics+of+ribosome+subunit+association+studied+by+mixing-spraying+time-resolved+cryogenic+electron+microscopy%5BTitle%5D)+AND+%22Structure%22%5BJournal%5D)">Structural dynamics of ribosome subunit association studied by mixing-spraying time-resolved cryogenic electron microscopy.</a> Structure. 23 (6), 1097-1105 (2015).</li>
<li>Kaledhonkar, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Late+steps+in+bacterial+translation+initiation+visualized+using+time-resolved+cryo-EM%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Late steps in bacterial translation initiation visualized using time-resolved cryo-EM.</a> Nature. 570 (7761), 400-404 (2019).</li>
<li>Fu, Z., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+structural+basis+for+release-factor+activation+during+translation+termination+revealed+by+time-resolved+cryogenic+electron+microscopy%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">The structural basis for release-factor activation during translation termination revealed by time-resolved cryogenic electron microscopy.</a> Nature Communications. 10 (1), 2579(2019).</li>
<li>Mäeots, M. -E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Modular+microfluidics+enables+kinetic+insight+from+time-resolved+cryo-EM%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Modular microfluidics enables kinetic insight from time-resolved cryo-EM.</a> Nature Communications. 11 (1), 3465(2020).</li>
<li>Kaledhonkar, S., Fu, Z., White, H., Frank, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Time-resolved+cryo-electron+microscopy+using+a+microfluidic+chip%5BTitle%5D)+AND+%22Methods+in+Molecular+Biology%22%5BJournal%5D)">Time-resolved cryo-electron microscopy using a microfluidic chip.</a> Methods in Molecular Biology. 1764, 59-71 (2018).</li>
<li>Jain, T., Sheehan, P., Crum, J., Carragher, B., Potter, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Spotiton%3A+a+prototype+for+an+integrated+inkjet+dispense+and+vitrification+system+for+cryo-TEM%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">Spotiton: a prototype for an integrated inkjet dispense and vitrification system for cryo-TEM.</a> Journal of Structural Biology. 179 (1), 68-75 (2012).</li>
<li>Razinkov, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+new+method+for+vitrifying+samples+for+cryoEM%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">A new method for vitrifying samples for cryoEM.</a> Journal of Structural Biology. 195 (2), 190-198 (2016).</li>
<li>Dandey, V. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Spotiton%3A+New+features+and+applications%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">Spotiton: New features and applications.</a> Journal of Structural Biology. 202 (2), 161-169 (2018).</li>
<li>Dandey, V. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Time-resolved+cryo-EM+using+Spotiton%5BTitle%5D)+AND+%22Nature+Methods%22%5BJournal%5D)">Time-resolved cryo-EM using Spotiton.</a> Nature Methods. 17, 897-900 (2020).</li>
<li>Wei, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Optimizing+%22self-wicking%22+nanowire+grids%5BTitle%5D)+AND+%22Journal+of+Structural+Biology%22%5BJournal%5D)">Optimizing "self-wicking" nanowire grids.</a> Journal of Structural Biology. 202 (2), 170-174 (2018).</li>
<li>Ravelli, R. B. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cryo-EM+structures+from+sub-nl+volumes+using+pin-printing+and+jet+vitrification%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Cryo-EM structures from sub-nl volumes using pin-printing and jet vitrification.</a> Nature Communications. 11 (1), 2563(2020).</li>
</ol>

B.C./C.S.P. have a commercial relationship with SPT Labtech, a company that produces a commercially available instrument, chameleon, that is based on the Spotiton prototype.

This protocol outlines the use of the Spotiton robotic system to prepare grids for cryo-EM imaging that carry two samples, generally a protein of interest and an activating ligand, that have been mixed for 90-500 ms. Although the workflow is straightforward, there are a few considerations the user must keep in mind to ensure a productive grid-making session. First, it is not uncommon for one of the piezoelectric dispenser tips to become clogged or blocked preventing it from firing. Such a failure will result in a liquid stripe, seen on image captures from the upper or lower camera, which is visibly narrower than that seen after both tips fired. A blockage could result from an air bubble lodging in the tip, disrupting droplet formation, or from protein sample that has dried out and sealed the narrow tip orifice. Although the aspirated sample is lost in the process, both problems can be resolved by an ultrasonic wash of the tips and re-aspiration of the sample. To prevent subsequent clogging and sample waste, it is crucial to thoroughly prime (flush) the fluid lines prior to sample aspiration and to maintain a high and consistent humidity level within the chamber and the shroud. Additionally, a protein sample with a particularly high concentration can affect tip firing despite well-maintained humidity. Although increasing the firing amplitude on the Inspect tab&#160;can partially compensate for weak firing due to high protein concentration, diluting the sample at least 1:2 will improve tip firing and avoid clogging.
Second, it can be difficult to achieve the ideal wicking speed that is needed to generate ice of optimal thickness for the targeted mixing duration. Generally, faster mixing times will require faster wicking, slower mixing times require slower wicking. For the ideally wicked grid, a liquid stripe is clearly discernable in the upper camera image, while in the lower camera, only a very slight thickening of the grid bars in the location of the stripe remains visible. Slow wicking, indicated by a dark stripe on the lower camera image, generally leaves ice that is too thick for imaging. Absence of a stripe on either image indicates fast wicking that may have left no water in the holes (Figure S5). Several factors such as nanowire density, plasma cleaning settings and duration, and time of exposure to and set level of chamber humidity can affect wicking speed. Poor (slow) wicking may be the result of a sparse coating of nanowires on the grid. By slightly diluting and increasing the time of exposure to the nanowire solution16, the density and coverage of nanowires on the grid bars will increase, facilitating faster wicking. If the nanowire density is sufficient, increasing the wattage setting or duration of plasma cleaning will also improve wicking. The settings recommended here are relatively low power and long duration, but may be changed if needed.
However, if both the specific batch of grids and plasma cleaning settings have worked well in a prior session, slow wicking performance may arise from excessive exposure of the nanowires to high humidity within the chamber, leading to their saturation with moisture and reduced liquid-holding capacity. The grid-saturating effect of chamber humidity can be reduced by either minimizing the elapsed time between tweezer mounting and grid plunging or decreasing the system humidity level prior to a plunge. It should be noted, however, that the latter brings the associated risk that the tip loaded with protein sample will clog. To offset this risk, holding the tips in the shroud where a high humidity level is maintained, can increase the allowable amount of time to mount tweezers holding a new grid. Finally, it should be noted that a reduced plunge time (achieved by increasing grid acceleration and/or maximum velocity) can result in grids with thinner ice without actually changing the wicking characteristics of the grid. However, as the mixing time of the two samples will also be reduced, plunge time is not a factor that is typically changed to address slow wicking. To address wicking that is too fast, resulting in ice that is either too thin or absent in the grid holes, the opposite of the measures outlined above can be taken.
Spotiton presents certain advantages and disadvantages when compared to other techniques developed for subsecond time-resolved studies. As the mixed sample stripe contains only 2-4 nL of liquid from each dispenser, a single 3 &#181;L aliquot of each sample is sufficient to prepare many grids-a key advantage when sample is limited. Additionally, observation of sample deposition using integrated cameras, although not entirely unique to Spotiton17, is not a feature of other mixing devices and allows plunged grids to be subjected to a crude pass/fail evaluation, greatly reducing screening time. One key disadvantage of the system is a minimum mixing time of 90 ms, restricted by the physical limitations of the mechanical components, that puts interrogation of faster biological reactions out of reach. By comparison, times less than 10 ms are routinely achieved on existing microfluidic systems. On the Spotiton-based, commercially available chameleon system, design and construction improvements have reduced the minimum plunge time to 54 ms and raise the possibility that addition of a second dispenser could allow faster mixing times than Spotiton can currently offer.
To date, a range of experiments has been conducted to investigate early, short-lived molecular states using Spotiton, including assembly of the 70S ribosome, calcium-triggered conformational changes in a transmembrane ion channel, and constriction of dynamin in response to GTP hydrolysis15. Since the publication of these results, several changes have been incorporated to the system to enhance the throughput, reproducibility, and reporting of Spotiton grid-making sessions. These include, among others, the dual-zone, automatic humidity-monitoring and control system, the experiment viewer feature, the side-by-side tip inspection feature, and several minor upgrades to the UI. The upgraded system will better support future two-sample mixing experiments similar to those previously reported as well as rapid binding assays such as between a small-molecule therapeutic and its protein target or even antibody-antigen complex formation. While current and future time-resolved experiments involving two interacting partners will certainly continue, the addition of a third piezoelectric dispenser and associated hardware could further broaden the range of possible experiments. For example, initial deposition of a detergent followed by the protein of interest, followed by the interacting or activating ligand could remove any potential negative impact of extended exposure to the detergent, often necessary to prevent common suboptimal imaging outcomes such as preferred orientation. In light of both the already published work and potential future applications, Spotiton represents an important tool for the cryo-EM community to facilitate the conduct of subsecond time-resolved studies.

The potential for cryo-EM to capture and reveal transient conformational states of proteins on the sub-second timescale (time-resolved cryo-EM) has been realized by several groups, beginning with Berriman and Unwin1 whose technique built upon the standard plunge freezing method developed by Dubochet for preparing cryo-EM grids2. They added an atomizer just above the cryogen cup that used compressed nitrogen gas to spray a fine mist of a second sample onto a plunging EM grid containing a first sample that had been applied and blotted to a thin aqueous layer. Although this system could achieve mixing times as low as 1 ms, it still demanded manual blotting of the first sample by the user-a technically challenging task-and a relatively high volume of the second sample. Moreover, in practice, it was difficult to know where mixing of the two samples had occurred, requiring the use of ferritin nanoparticles as a fiducial marker in the mixed sample. Subsequent efforts by Howard White and co-workers improved control and reproducibility of this spraying-mixing approach by incorporating computer control of the blotting and spraying steps3,4. To address how well and where the samples mix, the same group5 and others6,7,8,9,10 have moved to a mixing-spraying approach11 in which two samples are mixed either in narrow capillary tubes under the pressure of syringe pumps or in microfabricated, microfluidic chips driven by nitrogen gas. These premixing systems not only ensure complete mixing, but also enable the fine-tuning of mixing times to increase the resolution of time-resolved studies.
The introduction of piezoelectric dispensers as an alternate way to apply sample to EM grids in the Spotiton system enabled both the precise targeting of sample deposition and the requirement of a much smaller sample volume to make a grid12. Later, the use of nanowire grids and en route sample application (&#34;on-the-fly&#34; spotting) removed the need for a blotting step and reduced application-to-vitrification times13,14. For the new approach to time-resolved cryo-EM described here, a second dispenser along with the necessary control hardware and software upgrades were added to the Spotiton system to enable the delivery of a second sample onto a moving nanowire grid almost immediately after deposition of the first15. The two overlapping samples mix on the grid as they are wicked by the nanowires into a thin aqueous layer prior to vitrification. Mixing times as low as 90 ms can be achieved. This protocol intends to provide practical information as to how to conduct time-resolved experiments using piezoelectric dispensing and nanowire grids. Additionally, as the hardware and software are being modified to enhance ease of use, consistency, and throughput, this protocol also serves as an up-to-date description of the previously reported method15.

<table><tbody><tr><td>Buffer</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>cryogenic grid storage boxes</td><td>EMS (Hatfield, PA; USA)</td><td>N/A</td><td></td></tr><tr><td>Cu/Rh 300 mesh EM grids</td><td>EMS (Hatfield, PA; USA)</td><td>EMS300-Cu-Rh</td><td>treated to coat with nanowires as previously described (Wei et al, 2018)</td></tr><tr><td>ethane gas</td><td>TW Smith Corp. (Brooklyn, NY; USA)</td><td>N/A</td><td></td></tr><tr><td>Grid-handling forceps</td><td>EMS (Hatfield, PA; USA)</td><td>78320-5B</td><td></td></tr><tr><td>liquid nitrogen</td><td>Airgas, Inc. (Radnor, PA; USA)</td><td>N/A</td><td></td></tr><tr><td>liquid nitrogen reservoir with brass ethane cup (from FEI Vitrobot)</td><td>ThermoFisher Scientific (Waltham, MA; USA)</td><td></td><td></td></tr><tr><td>Picosystem</td><td>Hydro System and Supplies (Durham, NC; USA)</td><td>N/A</td><td>water purification system</td></tr><tr><td>Protein/other sample</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Solarus 950 plasma cleaner</td><td>Gatan, Inc. (Pleasanton, CA; USA)</td><td>N/A</td><td></td></tr></tbody></table>

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.

Figure 4&#160;shows images of grids prepared during a single time-resolved Spotiton session by mixing RNA polymerase and a 105 bp DNA oligomer carrying a promoter sequence for 150 ms prior to vitrification15. Seen in the figure are images taken by the two high-speed cameras of six grids at two timepoints following sample application. These cameras, unique among plunge-style freezers, allow the user to immediately decide whether to keep or discard a grid based on the observed extent of wicking by the nanowires and the likelihood the liquid samples were drawn into an aqueous layer thin enough for EM imaging. Although a single grid can provide sufficient ice for a complete dataset, once optimal conditions have been achieved, several grids are prepared to keep on hand in the event one or more is lost or contaminated. Of the six grids prepared in this session, the image captures show only one with suboptimal wicking (Figure 4F).
The grids shown were prepared (steps 2.5.1 through 2.5.8) in succession over a period of 40 min after an hour to ready the samples and the machine as described in the protocol (steps 1.1.1 through 2.4.6). Of the six grids, two were used for data collection, and the rest were saved for later analysis if needed. The pattern of ice on a vitrified grid (Figure 5C) matches closely the pattern of deposited liquid seen in the upper camera image (Figure 5B). Effective wicking of the mixed samples, made apparent by the lack of a visible liquid stripe in the lower camera image (Figure 5A), occurs along the nanowire-covered grid bars, and sample rarely overflows into squares adjacent to those in which it landed. Within ice-filled squares, the ice is typically thickest within holes at the center of the square and becomes thinner in holes closer to the grid bars (Figure 5E). Often holes immediately adjacent to the grid bars are empty due to proximity to the nanowires (Figure 5F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62271/62271fig01.jpg" /> 
Figure 1: The time-resolved Spotiton system. 1. Operator&#39;s workstation; 2. Environmental chamber; 3. Nitrogen supply; 4. Ethane supply 5. Backlight control for upper plunge path camera 6. Piezoelectric dispenser controllers; 7. Syringe pumps; 8. Vacuum pump; 9. Wash water supply and waste bottles. <a href="https://www.jove.com/files/ftp_upload/62271/62271fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62271/62271fig02.jpg" /> 
Figure 2: Spotiton software user interface. (A) Desktop icon and splash screen. (B) The main UI is composed of six areas: 1. display area for upper plunge path (&#34;upper&#34;) and tip inspection cameras; 2. display area for lower plunge path (&#34;lower&#34;) camera; 3. Playback area for tip inspection videos; 4. Multifunction tab area; 5. Live humidity monitor; 6. Live system logfile. Abbreviation: UI = user interface. <a href="https://www.jove.com/files/ftp_upload/62271/62271fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62271/62271fig03.jpg" /> 
Figure 3: Interior view of Spotiton chamber (robots in SafePosition). 1. Grid robot (red); 2. Dispenser robot (yellow); 3. Upper plunge path camera (pink); 4. Lower plunge path camera (light blue); 5. Tip inspection camera (orange); 6. Humidity shroud (green); 7. sample tray; 8. Nebulizer assembly (dark blue); 9. System reservoir and water lines. <a href="https://www.jove.com/files/ftp_upload/62271/62271fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62271/62271fig04.jpg" /> 
Figure 4: Representative system camera images from a time-resolved Spotiton grid making session. (A-F) Upper (left) and lower (right) plunge path camera images of six grids onto which RNA polymerase and a promoter DNA sequence were applied using Spotiton. Scale bar = 500 &#181;m. <a href="https://www.jove.com/files/ftp_upload/62271/62271fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62271/62271fig05.jpg" /> 
Figure 5: Pattern of ice deposition on a Spotiton-prepared grid. Portions of the (A) lower and (B) upper camera images and (C) atlas of the grid shown in Figure 4F. Approximate locations of ice resulting from sample deposition and mixing are colored lavender. Areas within the square marked with a yellow arrowhead are imaged in (E-G). The region shown in (E) is boxed in yellow in (D). Representative square (E), hole (F), and (G) exposure images collected from the grid shown in (A-D). The boxed areas in the square and hole images correspond to the hole and exposure images, respectively. Scale bars = 100 &#181;m (A-D), 5 &#181;m (E), 2 &#181;m (F), 100 nm (G). <a href="https://www.jove.com/files/ftp_upload/62271/62271fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure S1: Inspection of tip firing. A live view of Tip 2 firing is seen in the upper camera monitor on the left of the User Interface, while a recording made previously of Tip 1 firing plays on a loop for comparison in the video playback area on the right.&#160;<a href="https://www.jove.com/files/ftp_upload/62271/Figure S1.jpg" target="_blank">Please click here download this File.</a>
Figure S2: Ultrasonic cleaning of dispenser tips. An air bubble in the (A) tip will disrupt droplet formation and prevent tip firing. (B) Tips immersed in water at the ultrasonic cleaning station to remove an air bubble or clear dried protein blocking the tip orifice.&#160;<a href="https://www.jove.com/files/ftp_upload/62271/Figure S2.jpg" target="_blank">Please click here download this File.</a>
Figure S3: Loading grid in tweezers. (A) A grid placed nanowire side up on the edge of the grid block. (B) A grid positioned correctly in the self-closing tweezers.&#160;<a href="https://www.jove.com/files/ftp_upload/62271/Figure S3.jpg" target="_blank">Please click here download this File.</a>
Figure S4: Humidity tracker. The percent relative humidity in the chamber (dark blue) and shroud (light blue) as recorded during a typical grid-making session. The times of grid plunges (green squares) are plotted on the graph.&#160;<a href="https://www.jove.com/files/ftp_upload/62271/Figure S4.jpg" target="_blank">Please click here download this File.</a>
Figure S5: Wicking on nanowire grids during a plunge. Representative upper (A, C, E) and lower (B, D, F) plunge path camera images of wicking on nanowire grids that is too slow (A, B), ideal (C, D), and too fast (E, F). A slight thickening (white arrowheads) indicates grid bars with nanowires that have been saturated by sample. Squares in these regions typically contain ice of appropriate thickness for electron microscopic imaging. Scale bar = 500 &#181;m.&#160;<a href="https://www.jove.com/files/ftp_upload/62271/Figure S5.jpg" target="_blank">Please click here download this File.</a>

Watch this Scientific Journal Video about Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton at JoVE.com

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

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.

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental ...

cryo-electron-microscopic-grid-preparation-for-time-resolved-studies

Research

JoVE Journal

Biology

3.6K Views.  Simons Electron Microscopy Center, New York Structural Biology Center. 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.

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

Fast Grid Preparation for Time-Resolved Cryo-Electron Microscopy

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

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

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

Biochemistry

Using Tomoauto: A Protocol for High-throughput Automated Cryo-electron Tomography

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native context at a molecular level. The technique involves initially preserving the sample in its native state by rapidly freezing the specimen in vitreous ice, then collecting a series of micrographs from different angles at high magnification, and finally computationally reconstructing a 3-D density map. The frozen-hydrated specimen is extremely sensitive to the electron beam and so micrographs are collected at very low electron doses to limit the radiation damage. As a result, the raw cryo-tomogram has a very low signal to noise ratio characterized by an intrinsically noisy image. To better visualize subjects of interest, conventional imaging analysis and sub-tomogram averaging in which sub-tomograms of the subject are extracted from the initial tomogram and aligned and averaged are utilized to improve both contrast and resolution. Large datasets of tilt-series are essential to understanding and resolving the complexes at different states, conditions, or mutations as well as obtaining a large enough collection of sub-tomograms for averaging and classification. Collecting and processing this data can be a major obstacle preventing further analysis. Here we describe a high-throughput cryo-ET protocol based on a computer-controlled 300kV cryo-electron microscope, a direct detection device (DDD) camera and a highly effective, semi-automated image-processing pipeline software wrapper library tomoauto developed in-house. This protocol has been effectively utilized to visualize the intact type III secretion system (T3SS) in Shigella flexneri minicells. It can be applicable to any project suitable for cryo-ET.

We present a protocol on how to utilize high-throughput cryo-electron tomography to determine high resolution in situ structures of molecular machines. The protocol permits large amounts of data to be processed, avoids common bottlenecks and reduces resource downtime, allowing the user to focus on important biological questions.

Cryo-electron tomography (Cryo-ET) is a powerful three-dimensional (3-D) imaging technique for visualizing macromolecular complexes in their native ...

Bioengineering

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 ...

Preparation of Nucleosome Core Particles Complexed with DNA Repair Factors for Cryo-Electron Microscopy Structural Determination

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of chromatin, show most DNA repair enzymes remove DNA damage at reduced rates as compared to free DNA. The molecular details on how base excision repair (BER) enzymes recognize and remove DNA damage in nucleosomes have not been elucidated. However, biochemical BER data of nucleosomal substrates suggest the nucleosome presents different structural barriers dependent on the location of the DNA lesion and the enzyme. This indicates the mechanisms employed by these enzymes to remove DNA damage in free DNA may be different than those employed in nucleosomes. Given that the majority of genomic DNA is assembled into nucleosomes, structural information of these complexes is needed. To date, the scientific community lacks detailed protocols to perform technically feasible structural studies of these complexes. Here, we provide two methods to prepare a complex of two genetically fused BER enzymes (Polymerase &#946; and AP Endonuclease1) bound to a single-nucleotide gap near the entry-exit of the nucleosome for cryo-electron microscopy (cryo-EM) structural determination. Both methods of sample preparation are compatible for vitrifying quality grids via plunge freezing. This protocol can be used as a starting point to prepare other nucleosomal complexes with different BER factors, pioneer transcription factors, and chromatin-modifying enzymes.

This protocol outlines in detail the preparation of nucleosomal complexes using two methods of sample preparation for freezing TEM grids.

DNA repair in the context of chromatin is poorly understood. Biochemical studies using nucleosome core particles, the fundamental repeating unit of ...

Accelerating Macromolecular Structural Analysis Using Smart Leginon Autoscreen

Cryo-Electron Microscopy Screening Automation Across Multiple Grids Using Smart Leginon

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular protein complexes to near-atomic resolution. The general workflow of the entire cryoEM pipeline involves iterating between sample preparation, cryoEM grid preparation, and sample/grid screening before moving on to high-resolution data collection. Iterating between sample/grid preparation and screening is typically a major bottleneck for researchers, as every iterative experiment must optimize for sample concentration, buffer conditions, grid material, grid hole size, ice thickness, and protein particle behavior in the ice, amongst other variables. Furthermore, once these variables are satisfactorily determined, grids prepared under identical conditions vary widely in whether they are ready for data collection, so additional screening sessions prior to selecting optimal grids for high-resolution data collection are recommended. This sample/grid preparation and screening process often consumes several dozen grids and days of operator time at the microscope. Furthermore, the screening process is limited to operator/microscope availability and microscope accessibility. Here, we demonstrate how to use Leginon and Smart Leginon Autoscreen to automate the majority of cryoEM grid screening. Autoscreen combines machine learning, computer vision algorithms, and microscope-handling algorithms to remove the need for constant manual operator input. Autoscreen can autonomously load and image grids with multi-scale imaging using an automated specimen-exchange cassette system, resulting in unattended grid screening for an entire cassette. As a result, operator time for screening 12 grids may be reduced to ~10 min with Autoscreen compared to ~6 h using previous methods which are hampered by their inability to account for high variability between grids. This protocol first introduces basic Leginon setup and functionality, then demonstrates Autoscreen functionality step-by-step from the creation of a template session to the end of a 12-grid automated screening session.

Author Spotlight: Enhancing Cryo-Electron Microscopy by Automated Data Collection and Analysis Techniques

Cryo-electron microscopy (cryoEM) multi-grid screening is often a tedious process that demands hours of attention. This protocol shows how to set up a standard Leginon collection and Smart Leginon Autoscreen to automate this process. This protocol can be applied to the majority of cryoEM holey foil grids.

Advancements in cryo-electron microscopy (cryoEM) techniques over the past decade have allowed structural biologists to routinely resolve macromolecular ...

Preparation of Electron Microscopy Grids for In Situ Cellular Cryotomography

Sample Preparation for In Situ Cryotomography of Mammalian Cells

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly relevant to cellular biology and virology. The potential of combining cryotomography with other microscopy modalities makes it a perfect technique for integrative and correlative imaging. However, sample preparation for in situ cellular tomography is not straightforward, as cells do not readily attach and stretch over the electron microscopy grid. Additionally, the grids themselves are fragile and can break if handled too forcefully, resulting in the loss of imageable areas. The geometry of tissue culture dishes can also pose a challenge when manipulating the grids with tweezers. Here, we describe the tips and tricks to overcome these (and other) challenges and prepare good-quality samples for in situ cellular cryotomography and correlative imaging of adherent mammalian cells. With continued advances in cryomicroscopy technology, this technique holds enormous promise for advancing our understanding of complex biological systems.

Author Spotlight: Optimizing Grid Preparation for Enhanced Cryoelectron Tomography 

This method provides an accessible and flexible protocol for the preparation of electron microscopy (EM) grids for in situ cellular cryotomography and correlative light and electron microscopy (CLEM).

In situ cellular cryotomography is a powerful technique for studying complex objects in their native frozen-hydrated cellular context, making it highly ...

Fabrication of Streptavidin Affinity Grids for Cryo-EM Structural Studies

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

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

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

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

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

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 ...

A 3D Cartographic Description of the Cell by Cryo Soft X-ray Tomography

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these techniques, cryo soft X-ray tomography (SXT) allows imaging whole cryo-preserved cells in the water window X-ray energy range (284-543 eV), in which carbon structures have intrinsically higher absorption than water, allowing the 3D reconstruction of the linear absorption coefficient of the material contained in each voxel. Quantitative structural information at the level of whole cells up to 10 &#181;m thick is then achievable this way, with high throughput and spatial resolution down to 25-30 nm half-pitch. Cryo-SXT has proven itself relevant to current biomedical research, providing 3D information on cellular infection processes (virus, bacteria, or parasites), morphological changes due to diseases (such as recessive genetic diseases) and helping us understand drug action at the cellular level, or locating specific structures in the 3D cellular environment. In addition, by taking advantage of the tunable wavelength at synchrotron facilities, spectro-microscopy or its 3D counterpart, spectro-tomography, can also be used to image and quantify specific elements in the cell, such as calcium in biomineralization processes. Cryo-SXT provides complementary information to other biological imaging techniques such as electron microscopy, X-ray fluorescence or visible light fluorescence, and is generally used as a partner method for 2D or 3D correlative imaging at cryogenic conditions in order to link function, location, and morphology.

Here, a protocol describing the sample preparation and data collection steps required in cryo soft X-ray tomography (SXT) to image the ultrastructure of whole cryo-preserved cells at a resolution of 25 nm half pitch, is presented.

Imaging techniques are fundamental in order to understand cell organization and machinery in biological research and the related fields. Among these ...