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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+transient+structures+by+cryo-microscopy+combined+with+rapid+mixing+of+spray+droplets.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vitrification+of+pure+water+for+electron+microscopy.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+computer-controlled+spraying-freezing+apparatus+for+millisecond+time-resolution+electron+cryomicroscopy.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+second+generation+apparatus+for+time-resolved+electron+cryo-microscopy+using+stepper+motors+and+electrospray.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+cryo-EM+grid+preparation+device+for+time-resolved+structural+studies.">A cryo-EM grid preparation device for time-resolved structural studies.</a> IUCrJ. 6, 1024-1031 (2019).</li><li>Lu, Z., 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=Gas-assisted+annular+microsprayer+for+sample+preparation+for+time-resolved+cryo-electron+microscopy.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+dynamics+of+ribosome+subunit+association+studied+by+mixing-spraying+time-resolved+cryogenic+electron+microscopy.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Late+steps+in+bacterial+translation+initiation+visualized+using+time-resolved+cryo-EM.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+structural+basis+for+release-factor+activation+during+translation+termination+revealed+by+time-resolved+cryogenic+electron+microscopy.">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>M&#228;eots, M. -. E., 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=Modular+microfluidics+enables+kinetic+insight+from+time-resolved+cryo-EM.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-resolved+cryo-electron+microscopy+using+a+microfluidic+chip.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spotiton:+a+prototype+for+an+integrated+inkjet+dispense+and+vitrification+system+for+cryo-TEM.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+for+vitrifying+samples+for+cryoEM.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spotiton:+New+features+and+applications.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-resolved+cryo-EM+using+Spotiton.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimizing+"self-wicking"+nanowire+grids.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryo-EM+structures+from+sub-nl+volumes+using+pin-printing+and+jet+vitrification.">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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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

Nanyang Technological University

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

Preparation of Rat Brain Aggregate Cultures for Neuron and Glia Development Studies

An in vitro system that recapitulates the development and differentiation of progenitors into mature neurons and glia in the central nervous system (CNS) would provide a powerful platform for neuroscientists to investigate axo-glial interactions, properties and differentiation of multipotent progenitors, and progression of oligodendroglial lineage cells at the cellular and molecular level.  We describe here a CNS aggregate culture system from embryonic rat forebrains, which can be maintained in a serum-free medium up to 3-4 weeks and is used in our laboratory as a model to study neuron-glia interaction and CNS myelination.  This video clip will demonstrate how to isolate and grow these CNS aggregate cultures from E16 rat brain.  Furthermore, from the same brain dissection, highly enriched regular dissociated neuronal cultures can be readily obtained and used for various studies on CNS neurons or used for co-cultures with other cells. 

A protocols for an embryonic rat brain aggregate culture system is described. Multipotent progenitors in the aggregates can develop and differentiate into neurons, astrocytes and oligodendrocytes.

An in vitro system that recapitulates the development and differentiation of progenitors into mature neurons and glia in the central nervous system (CNS) ...

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System 

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers. Genetic studies in mammalian systems have been limited due to the lack of readily available tools including defined mutant genetic cell lines, regulatory expression systems, and appropriate selectable markers. To circumvent these difficulties, model genetic systems in lower eukaryotes have become an attractive choice for the study of functionally conserved DNA repair proteins and pathways. We have developed a model yeast system to study the poorly defined genetic functions of the Werner syndrome helicase-nuclease (WRN) in nucleic acid metabolism. Cellular phenotypes associated with defined genetic mutant backgrounds can be investigated to clarify the cellular and molecular functions of WRN through its catalytic activities and protein interactions. The human WRN gene and associated variants, cloned into DNA plasmids for expression in yeast, can be placed under the control of a regulatory plasmid element. The expression construct can then be transformed into the appropriate yeast mutant background, and genetic function assayed by a variety of methodologies. Using this approach, we determined that WRN, like its related RecQ family members BLM and Sgs1, operates in a Top3-dependent pathway that is likely to be important for genomic stability. This is described in our recent publication [1] at <a href="http://www.impactaging.com">www.impactaging.com</a>. Detailed methods of specific assays for genetic complementation studies in yeast are provided in this paper.

Genetic Studies of Human DNA Repair Proteins Using Yeast as a Model System

Genetic studies in yeast can be employed to investigate the molecular and cellular functions of human genes in cellular DNA metabolism. Methods are described for the genetic characterization of the human WRN gene product defective in the premature aging disorder Werner syndrome in functionally conserved pathways using yeast as a tractable model system.

Understanding the roles of human DNA repair proteins in genetic pathways is a formidable challenge to many researchers.  Genetic studies in mammalian ...

Preparation of Rat Tail Tendons for Biomechanical and Mechanobiological Studies

Rat tail tendons (RTTs) are a common biological model used in experimental in vitro studies in the fields of tendon physiology and tendinopathy. Working with those tissues is challenging because they are very fragile, and until now there was no rigorously detailed protocol for their isolation.
Faced with these challenges, we have developed methods and instruments to facilitate manipulation of RTTs and control tissue viability, sterility and integrity. This article describes the experimental procedures used to prepare RTTs for biomechanical and mechanobiological studies. Our work is divided into four main steps: extraction, cross-sectional area measurement, rinsing and loading into the bioreactor chamber.
At each step, all procedures, materials and manipulations are presented in detail so that they can be easily reproduced. Moreover, the specific instruments developed are presented: a manipulation plate used to segregate RTTs, an optic micrometer to position the tissue during the cross-sectional area measurement and an anchoring system to attach the RTTs onto a bioreactor.
Finally, we describe the results obtained after multiple tests to validate our methods. The viability, sterility and integrity evaluations demonstrate that our procedures are sufficiently rigorous for manipulations of fragile tissues such as rat tail tendons.

This article describes the experimental procedures used to prepare rat tail tendons for biomechanical and mechanobiological studies. Several features of the main steps in preparation are demonstrated, beginning with extraction, cross-sectional area measurement, rinsing and loading into the bioreactor chamber.

Rat tail tendons (RTTs) are a common biological model used in experimental in vitro studies in the fields of tendon physiology and tendinopathy. Working ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

High-throughput Saccharification Assay for Lignocellulosic Materials

Polysaccharides that make up plant lignocellulosic biomass can be broken down to produce a range of sugars that subsequently can be 
used in establishing a biorefinery. These raw materials would constitute a new industrial platform, which is both sustainable and carbon neutral, to replace 
the current dependency on fossil fuel. The recalcitrance to deconstruction observed in lignocellulosic materials is produced by several intrinsic properties 
of plant cell walls. Crystalline cellulose is embedded in matrix polysaccharides such as xylans and arabinoxylans, and the whole structure is encased by the 
phenolic polymer lignin, that is also difficult to digest 1. In order to improve the digestibility of plant materials we need to discover the main bottlenecks 
for the saccharification of cell walls and also screen mutant and breeding populations to evaluate the variability in saccharification 2. These tasks require 
a high throughput approach and here we present an analytical platform that can perform saccharification analysis in a 96-well plate format. This platform has 
been developed to allow the screening of lignocellulose digestibility of large populations from varied plant species. We have scaled down the reaction volumes 
for gentle pretreatment, partial enzymatic hydrolysis and sugar determination, to allow large numbers to be assessed rapidly in an automated system.
This automated platform works with milligram amounts of biomass, performing ball milling under controlled conditions to reduce the plant 
materials to a standardised particle size in a reproducible manner. Once the samples are ground, the automated formatting robot dispenses specified and recorded 
amounts of material into the corresponding wells of 96 deep well plate (Figure 1). Normally, we dispense the same material into 4 wells to have 4 replicates for 
analysis. Once the plates are filled with the plant material in the desired layout, they are manually moved to a liquid handling station (Figure 2). In this 
station the samples are subjected to a mild pretreatment with either dilute acid or alkaline and incubated at temperatures of up to 90&deg;C. The pretreatment solution 
is subsequently removed and the samples are rinsed with buffer to return them to a suitable pH for hydrolysis. The samples are then incubated with an enzyme
mixture for a variable length of time at 50&deg;C. An aliquot is taken from the hydrolyzate and the reducing sugars are automatically determined by the MBTH 
colorimetric method.

A simple, rapid method for determining the saccharification potential of large numbers of plant biomass samples is described. The automated platform for this analysis involves the preparation of the plant biomass for analysis in 96 well plates and the subsequent performance of pretreatment, hydrolysis and quantification of the sugars released.

Polysaccharides that make up plant lignocellulosic biomass can be broken down to produce a range of sugars that subsequently can be 
used in establishing ...

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 ...

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.

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.

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 ...

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 ...

Sample Preparation by 3D-Correlative Focused Ion Beam Milling for High-Resolution Cryo-Electron Tomography

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, frozen-hydrated state. However, cryo-ET requires that samples are thin enough to not scatter or block the incident electron beam. For thick cellular samples, this can be achieved by cryo-focused ion beam (FIB) milling. This protocol describes how to target specific cellular sites during FIB milling using a 3D-correlative approach, which combines three-dimensional fluorescence microscopy data with information from the FIB-scanning electron microscope. Using this technique, rare cellular events and structures can be targeted with high accuracy and visualized at molecular resolution using cryo-transmission electron microscopy (cryo-TEM).

Here, we present a pipeline for 3D-correlative focused ion beam milling on guiding the preparation of cellular samples for cryo-electron tomography. The 3D position of fluorescently tagged proteins of interest is first determined by cryo-fluorescence microscopy, and then targeted for milling. The protocol is suitable for mammalian, yeast, and bacterial cells.

Cryo-electron tomography (cryo-ET) has become the method of choice for investigating cellular ultrastructure and molecular complexes in their native, ...